Download Epitheliocystis in Lake Trout (Salvelinus namaycush)

Epitheliocystis in Lake Trout (Salvelinus namaycush)
by
Elena Contador
A Thesis
presented to
The University of Guelph
In partial fulfillment of requirements
for the degree of
Master of Science
Guelph, Ontario, Canada
© Elena Contador, January, 2013
ABSTRACT
EPITHELIOCYSTIS IN LAKE TROUT (SALVELINUS NAMAYCUSH )
Elena Contador
Advisor:
University of Guelph, 2013
Dr. John S. Lumsden
Epitheliocystis is a condition that has been recognised in a wide range of freshwater and
marine fish species. The infection is characterized by intracytoplasmic inclusions in gills
and less commonly, skin. Since 2004, molecular evidence has shown that epitheliocystis
is associated with Chlamydia-like organisms. Lake trout (Salvelinus namaycush) are
raised for release by the Ontario Ministry of Natural Resources at Blue Jay Creek,
Manitoulin Island, ON. This facility has experienced yearly epizootics of epitheliocystis
during winter. Affected lake trout have marked gill lesions including: severe branchial
epithelial cell necrosis, epithelial hyperplasia with thickening and blunting of lamellae
and occasionally small to large numbers of scattered epithelial cells containing 10-30 μm
bacterial inclusions. The use of PCR primers previously described to identify chlamydialike organisms has not produced consistent results with the gills of these LT with
prominent intraepithelial microcolonies. The use of universal bacterial primers to 16S
rDNA and laser capture microdissection in this thesis provide evidence that a novel βproteobacteria is more consistently associated with affected gills of LT and that this
organism may play an important role in these epizootics of EP.
ACKNOWLEDGEMENTS
I would like to express my gratitude to my supervisor, Dr. John Lumsden, whose
understanding, and patience were very important during all this process. I would like to
thank the other members of my committee, Dr. Salvatore Frasca, and Dr. Brandon Lillie
for their assistance in this research project. I must also acknowledge Paul Methner,
operations coordinator of the Blue Jay Creek Fish Culture Station (Manitoulin Island,
ON) without him the practical part of this project (sample collection and shipping) would
have been impossible. I thank my laboratory mates and friends in the Fish Pathology
Laboratory for teaching me English and for all the fun we have had.
I would like to thank my family; Roberto Contador, for all those nice conversations
which demonstrated the smart and open minded person he is and my mom Alicia Heredia
for potentiate my curiosity towards science during my childhood. I thank to my siblings
Roberto and Carito for always being there, listening or laughing with me. I would also
like to thank my husband and best friend, Pato, without whose love and support, I would
not have finished this thesis. Also I thanks to my friends outside the university, they have
helped to fill that space that is left when family is away.
This research would not have been possible without the financial assistance of NSERC,
OMNR and OMAFRA, the Department of Pathobiology at the University of Guelph and
the CONICYT Chilean Scholarship fund.
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TABLE OF CONTENTS
LIST OF LIST OF TABLES...……………………………………………………..…...viii
LIST OF LIST OF FIGURES……………………………………………………………ix
LIST OF ABBREVIATIONS…………………………………………………………….x
DECLARATION OF WORK PERFORMED……………………………………..…...xiii
GENERAL INTRODUCTION…………………………………………………...……....1
1. REVIEW OF LITERATURE..........................................................................................3
1.1. Epitheliocystis…………………………………………………………………….…..3
1.2. Histopathological findings…………………………………………………………....3
1.3. Electron microscopy (EM)……………………………………………………………5
1.4. Immunohistochemistry (IHC)…………………………………………………….......8
1.5. Polymerase chain reaction (PCR)…………………………………………………….9
1.6. In situ hybridization (ISH)…………………………………………………………..10
1.7. Agent………………………………………………..…………………………….....11
1.8. Order Chlamydiales and its association with EP in fish…………………………….12
1.9. CLOs identified in EP affected fish by antigenic and molecular methods……….....17
1.9.1. Candidatus Piscichlamydia salmonis (CPs)……………………………….……...17
1.9.2. Candidatus Clavochlamydia salmonicola (CCs)……………………………….....18
1.9.3. Neochlamydia spp…………………………………………….…………………...18
1.9.4. Candidatus Parilichlamydia carangidicola………………………………………..18
1.9.5. Candidatus Actinochlamydia clariae………………………………………….......19
1.9.6. Candidatus Renichlamydia lutjani………………………………..…………….....19
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1.10. Non-chlamydial agents associated with epitheliocystis…………………………....19
1.10.1. Candidatus Brachiomona cysticola (CBc)……………………….……………...19
1.10.2. Endozoicomonas elysicola (Ee)……………………………………….…………20
1.10.3. Rickettsia like organism (RLO)…………………………………….…………....20
1.10.4. Other agents……………………………………...................................................21
1.11. Transmission and pathogenesis …………………………………………………...22
1.12. Control and treatment…………………………………………………...………....26
1.13. Epitheliocystis in lake trout………………………………………………..……....27
1.14. Conclusion…………………………………..………………………………..........29
1.15. Rationale……………………………………..…………………………….............29
1.15.1. Purpose………………………………………………..………………………….30
1.15.2. Hypothesis & objectives……………………………………………....................30
2. MATERIALS AND METHODS……………………………………………………..33
2.1. Background……………………….…………………………………………...........33
2.2. Retrospective, prospective sample collection and tissue processing………………..33
2.3. Bacterial isolation…………….………………………………………….……….....36
2.4. Histopathology…….………………………………………………………………...36
2.4.1. Scoring of gill morphology………………………………………………………..37
2.5. DNA extraction…………………………..……………………………………….....39
2.5.1. Gill tissue……………………….…………………………....................................39
2.5.2. Bacterial isolates from gill tissue……….................................................................39
2.5.3. DNA extraction from formalin-fixed paraffin-embedded gill tissues…………….40
2.6. Polymerase chain reaction (PCR)……….…………………………………………..41
v
2.6.1. Primers……….………………………………………………………………........41
2.6.2. Amplification protocols……………………………………………………….......42
2.7. Electrophoresis and DNA purification………………………………………….…..48
2.8. DNA sequencing and analysis……………………………………….……...............48
2.9. Immunohistochemistry…………………………………..……………………….....49
2.10. Transmission electron microscopy…………………………………………….…..49
3. RESULTS…………………………………………………………………………….51
3.1. History and clinical findings…………………………………………….………….51
3.2. Bacterial isolation…………………………………………..…………………….....52
3.3. Histopathology…………………………………….………………………………..56
3.4. PCR and DNA sequencing analysis………………………………………………...69
3.4.1. Universal bacterial 16S rRNA gene primers……………………………………...69
3.4.2. 16rRNA Chlamydiales primers……………………………….…………………..75
3.4.3. EEDV primers................................................................................................….....80
3.5. Molecular and phylogenetic analyses……………………………………………….80
3.6. Immunohistochemistry…………..……………………………………………..…...84
3.7. Transmission electron microscopy………………………..…………………...........86
4. DISCUSSION.....................................................................................................……..93
5. CONCLUSIONS………………………………………… …………………............103
6. REFERENCES…………………………...………………. ……………….………..104
7. APPENDICE……………………………………………..………………….……....119
7.1. EP-PM Protocol ………………………………………..………………….……....119
7.2. Primers trouble shooting………………………………..…………………….…....123
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7.3. PCR protocols……………………………………………………………..……....124
7.4 Sequencing results from 2010 samples using 16S universal bacteria primers..........127
vii
LIST OF TABLES
Table 1.1 Epitheliocystis reported in farmed salmonids…………………………………28
Table 1.2: Epitheliocystis reported in wild salmonids…………………………………...28
Table 2.1 Lake trout epitheliocystis cases submitted to the Fish Pathology Laboratory
between 2007 and 2013…………………………………….............................................35
Table 2.2 Semiquantitative scores in lake trout gill lesions using light microscopy…….38
Table 2.3 List of primers used in this study………………………………………..…....46
Table 3.1 Sixteen bacterial isolates derived from fish with EP........................................54
Table 3.2 Lesion score results from tank R8 during 2013………………………............67
Table 3.3 Number of samples that have 90 % maximum identity with Candidatus
Brachiomona cysticola…………………………………………………………………..72
Table 3.4 Nucleotide similarities between sequences obtained by laser capture
microdissection and PCR from DNA directly extracted from fresh gill tissue of lake trout
with epitheliocystis……………………………………………........................................73
Table 3.5Sequences identified in lake trout during epitheliocystis outbreaks using 16S
Chlamydiales primers……………………………............................................................77
Table 3.6 Comparison between organisms identified by electron microscopy in previous
reports and the EPI organism described in the present study……………………….......87
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LIST OF FIGURES
Figure 1.1 Chlamydial developmental cycle………………………….………………....13
Figure.1.2 Streamlined representation of the order Chlamydiales……………………....16
Figure.2.1 Chlamydiales primer localization along the 16S rRNA gene of positive control
Candidatus Piscichlamydia salmonis………………………………………………........45
Figure 3.1 Pattern of daily mortality of LT during winter 2013 at BJC in tank R8…..…53
Figure 3.2 Histopathology of lake trout from 2006……………………………………...58
Figure 3.3 Histopathology of lake trout gills from 2013, group A……………….….…..59
Figure 3.4 Histopathology of lake trout gills from 2013, group B………………….…...60
Figure 3.5 Histopathology of lake trout gills from 2013, group B (2)………...................61
Figure 3.6 Histopathology of lake trout gills from 2013, group C……………….…...…62
Figure 3.7 Histopathology of lake trout gills from 2013, group D………………………63
Figure 3.8 Differential staining of lake trout gills from 2013, group D ………………...64
Figure 3.9 Gill lesions at four different times and mortality of Blue Jay Creek lake trout
from tank R8 during 2013…………………......................................................................68
Figure 3.10 Agarose gel (1 %) of 800 bp products generated from lake trout gills using
the PCR protocol UNIRX2 to detect bacterial 16S rDNA…………….………………...71
Figure 3.11 Tissue sections used for laser capture microdissection…………….….........74
Figure 3.12 Agarose gel (2 %) of 270 bp products generated from lake trout gills using
the PCR protocol CLORX4T……………………………………………….…………....79
Figure 3.13 Order Burkholderiales phylogenetic tree…………………………………....81
Figure 3.14 Order Chlamydiales phylogenetic tree……………………………………...83
ix
Figure 3.15 Immunohistochemistry of lake trout gills from 2013, group D …................85
Figure 3.16 Transmission electron microscopy of a lake trout gill from a fish affected by
epitheliocystis……………………………………………………………………………88
Figure 3.17 Transmission electron microscopy of lake trout gills affected by
epitheliocystis (1)………………………………………………………………………..89
Figure 3.18 Transmission electron microscopy of lake trout gill affected by
epitheliocystis (2)………………………………………………………………………..90
Figure 3.19 Transmission electron microscopy of lake trout gill affected by
epitheliocystis (3)………………………………………………………………………..91
Figure 3.20 Transmission electron microscopy of lake trout gill affected by
epitheliocystis (4)………………………………………………………………………..92
x
LIST OF ABBREVIATIONS
AS
Atlantic salmon
AC
Arctic charr
BA
blood agar
BFCG
Agar made of: blood, fetal bovine serum, cysteine and glucose
BLAST
basic local alignment tool
BGD
bacterial gill disease
BT
brown trout
CA
cytophaga agar
CBc
Candidatus Brachiomona cysticola
CCs
Candidatus Clavochlamydia salmonicola
CLO
Chlamydia-like organism(s)
CPs
Candidatus Piscichlamydia salmonis
d
day(s)
DNA
deoxyribonucleic acid
DEPI
deionized diethylpyrocarbonate
Ee
Endozoicomonas elysicola
EB
elementary body
EGC
eosinophilic granular cells
EM
electron microscopy
EP
epitheliocystis
EPI
epitheliocystis inclusion (s)
xi
FFPE
formalin-fixed paraffin-embedded
FPL
fish pathology laboratory
GM
goblet cells metaplasia
h
hour(s)
IB
intermediate body
H&E
hematoxylin and eosin
IHC
immunohistochemistry
ISH
in situ hybridization
KHV
koi herpes virus
LCM
laser capture microdissection
LM
light microscopy
LH
lamellar hyperplasia
LT
lake trout
min
minutes
MI
Maximum identity
N
necrosis
OMNR
Ontario Ministry of Natural Resources
PGI
proliferative gill inflammation
PCR
polymerase chain reaction
PS
pseudomonas agar
PVK
Pierce van der Kamp
RB
reticulate body
rDNA
ribosomal deoxynucleic acid
xii
RNA
ribonucleic acid
rRNA
ribosomal ribonucleic acid
RLO
Rickettsia-like organism (s)
RT
rainbow trout
SS
suspended solids
sec
seconds
TEM
transmission electron microscopy
TSA
trypticase soya agar
xiii
DECLARATION OF WORK PERFORMED
I declare that all work in this subject is my own with the exception of the following:
Transmission electron microscopy semi-thin sections and pictures by Paul Huber
Immunohistochemistry staining by Susan Lapos
xiv
GENERAL INTRODUCTION
Epitheliocystis (EP) is a world-wide condition that affects fish in fresh and salt water and
at a variety of temperatures. The term is used to describe a characteristic lesion, which
involves hypertrophied branchial epithelial cells with intracellular bacterial inclusions.
Over the last decade numerous advances on the identification of a causative EP agent
have been made, though none have been cultured yet. Amplification of partial or
complete segments, of the 16S rRNA gene by PCR and the localization of the obtained
sequences within lesions using in situ hybridization (ISH) have allowed the identification
and classification of different agents associated with this condition. Chlamydiales, ß- and
γ–proteobacteria have been identified as causes of intracellular inclusions; however, their
relationship with clinical signs and mortality remains unclear. EP has been associated
with mortality, but the presence of other infectious agents also associated with disease
and the lack of culture techniques have not allowed study of the role of EP agents in these
outbreaks. In lake trout (Salvelinus Namaycush) (LT), outbreaks associated with EP at
some Ontario Ministry of Natural Resources (OMNR) facilities have become an
impediment to the enhancement of this species. The histological lesions are fairly
consistent including prominent single-cell necrosis of leukocytes and epithelial cells,
thickening and blunting of lamellae. Antimicrobials used to date (Tribrissen® and
oxytetracycline) have not proven effective in limiting the impact of outbreaks at the
OMNR, Blue Jay Creeck (BJC) Fish Culture Station. Identification of the infectious
agent associated with mortalities in LT at BJC is the main aim of this study; this would
1
allow a better understanding of the pathogenesis, epidemiology and control of the
disease.
2
1. REVIEW OF LITERATURE
1.1. Epitheliocystis
Epitheliocystis (EP) is a bacterial condition with a worldwide distribution that has been
described in more than 50 freshwater and marine fish species (Paperna and Sabnai, 1980;
Lewia et al., 1992; Kent et al., 1998; Nowak and LaPatra, 2006; Aboei and Briyai, 2011).
The name of this condition was proposed by Hoffman to allow a differential diagnosis
between EP and lymphocystis in bluegill (Lepomis macrochirus). While EP was thought
to be caused by either a Rickettsia-like organism (RLO) or a Chlamydia-like organism
(CLO), lymphocystis was caused by a well-defined etiologic agent, an iridovirus
(Hoffman et al., 1969). Clinical disease has been attributed to respiratory insufficiency
secondary to branchial epithelial hyperplasia and excessive mucus production, although
evidence for this association remains circumstantial (Zimmer et al., 1984; Groff et al.,
1996).
1.2. Histopathological findings
EP affects the gill and skin; it has been defined as an “epithelial hyperplasia in response
to an infection, characterized by the presence of Gram-negative bacterial colonies
contained within epithelial cells. These bacteria are surrounded by an intracellular
vacuole that progressively enlarges and eventually results in the formation of spherical,
variably-sized cysts called inclusions” (Hoffman et al., 1969).
3
The morphology and branchial location of the inclusions or cysts may vary between host
fish species, age group and wild versus cultured fish. Characteristics such as cyst size,
agent morphology and host response (proliferative and non-proliferative) are some of the
variations noted between EP infections in different fishes (Paperna, 1978; Crespo et al.,
1999).
The reaction of the host gill tissue can range from little or no hyperplasia of the branchial
epithelium and inflammation of the interstitium (non-proliferative response) to a severe
response characterized by a marked lamellar epithelial hyperplasia and necrosis with mild
to marked inflammation (proliferative response) (Paperna, 1977; Crespo et al., 1999;
Draghi et al., 2004). The proliferative form of EP is similar to what has been termed
‘proliferative gill inflammation’ (PGI) in Atlantic salmon (Salmo salar) (AS), which
appears to be a multifactorial condition where CLO are commonly but not always
associated with disease (Kvellestad et al., 2005; Steinum et al., 2009). The diagnosis of
PGI is based on four criteria, all of them histopathological: 1) circulatory disturbances in
areas of pathological changes; 2) branchial epithelial hyperplasia; 3) infiltration of
inflammatory cells, and 4) the presence of necrosis in the affected tissue (Kvellestad et
al., 2005).
Light microscopy (LM) is a useful diagnostic tool when the inclusions are obvious,
although epithelial hyperplasia caused by an EP agent in the early stages of the disease
may not yet be accompanied by visible bacterial inclusions (Wolke et al., 1970; Nylud et
4
al., 1998). This can affect the accuracy of a histological diagnosis and generate false
negative reports. Histopathological evaluation of affected gills should be accompanied by
a complementary test in order to identify the organisms involved in EP outbreaks.
The lesions noted histologically are also markedly varied depending on the host species,
and this has provided support for the theory that there may be unique EP agents for each
fish species (Hoffman et al., 1969; Wolke et al., 1970; Paperna, 1977; Paperna et al.,
1981). However, molecular studies in salmonids, have shown that the same EP agent, e.g.
Candidatus Piscichlamydia salmonis (CPs), is able to infect AS, Artic charr (Salvelinus
alpinus) (AC), and brown trout (Salmo trutta) (BT) (Draghi et al., 2004; 2007; 2010;
Schmidt-Posthause et al., 2011). Lesion variation between fish species at least in some
salmonids is therefore more likely to be determined by the host response. Since there is
no study published to date that describes an infection over time, few conclusions can be
reached regarding the basis for lesion variation in one species let alone between species.
1.3. Electron microscopy (EM)
Since the infectious agent responsible for the ‘inclusions’ has not been cultured yet, the
initial studies on this disease were limited to histopathology and EM. Ultrastructurally,
the organisms within inclusions were described to be similar to RLO in striped bass
(Morone saxalitis), grey mullet (Liza ramada), golden grey mullet (Liza aurata), flathead
mullet (Mugil cephalus), American plaice (Hippoglosoides platessoides), silver perch
(Bidyanus bidyanus) and Tilapia spp., but were most similar to CLO in numerous other
5
species, including sea bream (Spaurus auratus), common carp (Cyprinus carpio), channel
catfish (Ictalurus punctatus), brown bullhead (Ameiurus nebulosus), greater amberjack
(Seriola dumerili), steelhead trout (Oncorhynchus mykiss), white sturgeon (Acipenser
transmontanus) and AS (Zachary and Paperna, 1977; Paperna et al., 1978; 1981; Paperna
and Sabnai, 1980; Molnar et al., 1981; Morrison and Shum, 1983; Paperna and Alves de
Matos, 1984; Rourke et al., 1984; Zimmer et al., 1984; Desser et al., 1988; Bradley et al.,
1988; Crespo et al., 1990; Grau and Crespo,1991; Padros and Crespo,1995; Groff et al.,
1986; Frances et al., 1997; Nylund et al., 1998; Szakolczai et al., 1999; Kim et al., 2004;
Agnetti et al., 2009).
Using EM, Wolke et al. (1970) described three common characteristics shared by CLOs
and the EP agent: intracytoplasmic location within a membrane-bound ‘inclusion’; the
particles within the inclusion have both a cell wall and a plasma membrane; and finally
the presence of four different developmental stages, including a giant body, an initial
body, an intermediate form and an elementary body. However, subsequent studies
reported structures that were not consistent with CLOs within infected cells (Wolke et al.,
197l; Zachary and Paperna, 1977). The microorganisms contained in these inclusions,
had pleomorphic developmental cycles and lacked some of the developmental stages of
CLO implying that they might be more similar to RLOs (Wolke et al., 1970; Zachary and
Paperna, 1977; Paperna et al., 1978; Molnar and Boros, 1981; Morrison and Shum, 1983;
Frances et al., 1997).
6
Subsequent morphological analyses using EM indicated potentially greater ultrastructural
variability in EP causative agents (Paperna et al., 1978; 1981; Rourke et al., 1984; Crespo
et al., 1999; Draghi et al., 2004). A study of affected epithelial cells of striped bass from
Chesapeake Bay (USA) showed that EP in this host was caused by rod-shaped
microorganisms structurally closer to rickettsia (Zachary and Paperna, 1977). In sea
bream and grey mullet there were several unusual forms in the same examined fish, such
as hyphae or mycelia-like structures as well as individual organisms connected by
cytoplasmic bridges; these are not characteristics consistent with either RLO or CLO
(Paperna et al., 1978). These studies lead to the possibility that either two kinds of EP
agents could be infecting the same individual at the same time, or infectious agents other
than members of the Chlamydiales may be associated with this condition (Paperna et al.,
1978; Crespo et al., 1999). These possibilities could only be proven by complementary
molecular and immunological tests (Nylund et al., 1998; Draghi et al., 2004; 2010).
Additional reasons for these structural variations in agents within the same affected fish,
the same fish species and among fish species may be related to the stage of growth and
maturity of the inclusion at the time of the sampling and/or environmental factors such as
salinity and temperature (Bradley et al., 1988; Nylund et al., 1998).
Dissimilar ultrastructural characteristics identified for the organisms present in these
inclusions in different fish species might also be attributed to inconsistent sample
processing. Several of these studies were performed on formalin-fixed tissue samples
(Hoffman et al., 1969; Wolke et al., 1970; Zimmer et al, 1984; Miyazaki et al., 1986;
7
Grau and Crespo, 1991). Improper tissue fixation for EM can cause dramatic alteration of
an bacterium’s morphology and the morphology of the membrane-bound vacuole and its
matrix. This is a highly likely cause of the variation described in the literature. This is
particularly critical with bacteria such as CLOs and RLOs, which are difficult to
differentiate structurally even with optimum fixation and staining (Anderson et al., 1965;
Wolke et al., 1970; Avakyan and Popov, 1984).
1.4. Immunohistochemistry (IHC)
Since the most commonly described agents associated with EP are CLOs, the IHC tests
performed in suspected samples to date have used antibodies to lipopolysaccharide
(LPS), which is common to all of the nine species found within the family
Chlamydiaceae. In most fish species affected by EP, there was no reactivity with
antibody to Chlamydia LPS antigen (Bradley et al., 1988; Langdon et al., 1991;
Anderson, 1992; Crespo et al., 1999; Meijer et al., 2006; Nowak and LaPatra, 2006;
Polkinghorne et al., 2010). It is possible that CLOs that do not belong to the
Chlamydiaceae family are the cause of EP in these fish. However, some studies have
successfully detected the Chlamydia LPS antigen within cytoplasmic inclusions,
specifically in cases affecting white sturgeon (Acipenser transmontanus), barramundi
(Latest calcarifer), silver perch, BT and AS, even though the CLO affecting these fish,
based on molecular phylogenetic analysis, do not belong to the family Chlamydiaceae. It
has been suggested that this might be explained by cross reactivity of the anti–LPS
antibodies with a related but as yet uncharacterized trisaccharide in the CLO (Groff et al.,
8
1996; Draghi et al., 2004; 2007; Meijer et al., 2006) and may also be explained by
intermittent expression of LPS antigen during CLO developmental cycle (Groff et al.,
1996).
1.5. Polymerase chain reaction (PCR)
PCR is currently a valuable tool in disease diagnosis; it has been utilised successfully for
taxonomy studies, generating phylogenetic and genotypic data that allows the
classification of organisms into specific groups by 16S rRNA gene amplification
(Fredricks and Relman, 1996; Woo et al., 2008; Austin, 2011). Increased knowledge
regarding the evolutionary relationships among groups is leading to a better
understanding of how the genome of pathogenic species or strains evolved from nonpathogenic ancestors and vice versa. This knowledge, along with phenotypic
characterizations, will help to clarify mechanisms of pathogenicity and strategies of host
adaptation (Weisburg et al., 1991; Palys et al., 1997; Daubin et al., 2002; Sudheesh et al.,
2012). As mentioned above, EP etiologic agents have not yet been cultured, which
complicates diagnosis and further research. However, the use of PCR primers to the 16S
rRNA gene in the Order Chlamydiales (Ossewaarde and Meijer, 1999; Everett et al.,
1999) have allowed identification of CPs in AS, BT and AC (Draghi et al., 2004; 2010;
Schmidt-Posthause et al., 2011), CRG18 (AY013394) in silver perch, CRG98
(AY013474) in barramundi CRG20 (AY013396) in the leafy sea dragon (Phycodorus
eques) (Meijer et al., 2006), UFC1 (FJ001668) in the leopard shark (Triakis semifasciata)
(Polkinghorne et al., 2010), Neochlamydia spp. (AY225593, AY225594) AC (Draghi et
9
al., 2007), Candidatus Clavochlamydia salmonicola (CCs) in AS and BT (Karlsen et al.,
2008; Schmidt-Posthause al., 2011) and recently Candidatus Parilichlamydia
carangidicola in yellowtail kingfish (Seriola lalandi) (Stride et al., 2013) and Candidatus
Actinochlamydia clariae in catfish (Clarias gariepinus). Two non-Chlamydiae bacteria,
Candidatus Brachiomona cysticola (CBc) and Endozoicomonas elysicola (Ee), were
identified from cases of EP in AS and cobia larvae (Rachycentron canadum) respectively,
using 16S rRNA universal bacterial primers (Toenshoff et al., 2012; Mitchell et al.,
2013; Mendoza et al., 2013).
PCR has proven to be extremely valuable when the disease agent in question is poorly
described, or cannot be isolated. However, chimera formation is influenced by multiple
factors including pairwise sequence identity between 16SrRNA genes, number of PCR
cycles and the relative abundance of gene-specific PCR templates. Chimeras are hybrid
products between multiple original sequences that can be falsely interpreted as novel
organisms, increasing the apparent diversity. In this case, post-amplification analyses of
the sequences can verify the quality of the sequence (Clarridge III, 2004, Hass et al.,
2011).
1.6. In situ hybridization (ISH)
Using Chlamydiales-specific and 16S rRNA universal primers, sequences were generated
from potential EP agents in infected gill tissues. Specific 16S rRNA oligoprobes or
riboprobes were then designed to localize CLO sequences within inclusions. Positive
10
labelling of CLO in inclusions has been demonstrated in silver perch, barramundi, AS
and AC among others (Draghi et al., 2004; 2007, 2010; Meijer et al., 2006). Positive
labelling of non-Chlamydiles agents CBs and Ec were demonstrated in inclusions using
fluorescent in situ hybridization (FISH) and ISH in AS and cobia larvae, respectively
(Toenshoff et al., 2012; Mendoza et al., 2013).
1.7. Agent
Bacteria found within eukaryotic cells can be either facultative or obligate intracellular
organisms. Inside an animal host cell, bacteria can reside in two different compartments.
They can be within a membrane-bound vacuole (called an inclusion in studies), which
may be derived from a phagosome formed during bacterial endocytosis, or they may
colonize the host cell cytosol (Goebel and Gross, 2001; Ochman and Moran, 2001).
Generally, bacteria capable of intracellular survival and growth have specific virulence
factors to recognize, invade, and multiply within eukaryotic cells (Zientz et al., 2004).
This intracellular location may allow exploitation of host metabolites in order to support
bacterial multiplication in a relatively safe host compartment in which it is partially
hidden from potent host defense mechanisms (Gross et al., 2003). Examples of bacteria
that are able to multiply inside an intracellular vacuole include: Salmonella enterica
serovar Typhimurium, Legionella pneumophila, Coxiella burnetii, Francisella tularensis,
Mycobacterium tuberculosis, and the obligate intracellular bacteria, Chlamydiae. Listeria
monocytogenes, Shigella flexneri, enteroinvasive Escherichia coli and some obligate
intracellular Rickettsia spp. are able to enter and replicate in the cytosol of mammalian
cells (Zientz et al., 2004). Obligate intracellular CLOs have been shown to be the putative
11
causative agents of EP (Draghi et al., 2004; 2007; 2010; Meijer et al., 2006; Karseln et
al., 2008; Mitchell et al., 2010; Schmidt-Posthause et al 2011; Straid et al., 2013; Steigen
et al., 2013). The facultative or obligate intracellular status of non-Chlamydiales EP agent
CBc is unknown. Ee is the intracellular EP agent in cobia larvae, and it has also been
isolated in agar media from the ornate leaf slug (Elysia ornata), demonstrating this
bacteria is a facultative intracellular agent. (Kurahashi and Yokota, 2007; Toenshoff et
al., 2012; Mendoza et al., 2013).
1.8. Order Chlamydiales and its association with EP in fish
Before their classification as bacteria, chlamydiae were previously categorized as
protozoa or viruses. Molecular evaluation of the 16S rRNA gene confirmed chlamydiae
to be a eubacteria with a very distant relationship to other eubacterial orders (Weisburg et
al., 1986). The organisms in this phylum share three characteristics: 1) small sized Gramnegative bacteria 2) obligate intracellular parasites of eukaryotic cells with a biphasic
development cycle, e.g. reticulate bodies (RB) and elementary bodies (EB) (Figure 1.1),
and 3). more than 80%16S rRNA sequence identity (Everett et al., 1999; Corsaro et al.,
2003; Corsaro and Greub, 2006).
12
Figure 1.1 Chlamydial developmental cycle (modified from Dr. Karin Everett,
http://chlamydiae.com/twiki/bin/view/Cell_Biology/GrowthCycle).
The term Chlamydia-like organism has been utilized to describe a bacterium that has an
obligate intracellular lifestyle, two developmental stages similar to Chlamydiaceae
members, but lack proof of the 16S rDNA sequence belonging to chlamydiales (Corsaro
et al., 2003; Corsaro and Greub, 2006; Horn, 2008). The prefix Candidatus applies when
there is evidence for the presence of all three characteristics, but the strains have not been
cultured or submitted to one of two reference culture collections e.g. CPs (Murray and
Stackebrandt, 1995; Euzeby and Tindal, 2004).
13
The concept of an ‘environmental Chlamydia’ is used to include those diverse
Chlamydiae isolated from environmental sources (Horn and Wagner, 2001; Horn et al.,
2004); however, a more general term such as ‘novel Chlamydia’ has been suggested to
include all those Chlamydiae that are not members of the family Chlamydiaceae (Corsaro
and Greub, 2006).
More than a decade ago, chlamydia were placed in their own order Chlamydiales, with
one family Chlamydiaceae containing a single genus, Chlamydia (Moulder et al., 1984).
Later the order Chlamydiales was then reorganized based on their 16S and 23S rDNA
sequence
identity
creating
four
families:
Chlamydiaceae,
Parachlamydiaceae,
Simkaniaceae and Waddliaceae (Everett et al. 1999). In the last decade, Candidatus
Rhabdochlamydiaceae (Kostanjsek et al., 2004), Candidatus Criblamydiaceae, (Thomas
et al., 2006) and Candidatus Parilichlamydiaceae (Stride et al., 2013), have been
suggested as novel families (Figure 1.1). The Chlamydiaceae, which previously
contained only the genus Chlamydia, was divided into two genera, Chlamydia and
Chlamydophila (Everett et al., 1999). All members of the Chlamydiaceae have 16S
rDNA genetic sequences with 90 % or greater nucleotide identity, while CLOs or
environmental chlamydias have 80-90 % nucleotide identity to members of the
Chlamydiaceae (Everett et al., 1999).
Despite the identification of CLOs in EP lesions by LM and EM in numerous fish since
1920s, it was not until 2002 that molecular evidence for the presence of Chlamydiae in
barramundi, leafy sea dragon and silver perch with EP was reported (Meijer and
14
Ossewaarde, 2002). Phylogenetic analysis showed that each of these EP agents clustered
with Chlamydia-like bacteria but were distinct from the Chlamydiaceae, although they
appear as new lineages within the order Chlamydiales (Meijer and Ossewaarde, 2002;
Meijer et al., 2006). These studies were the first to indicate the diversity of CLOs
associated with EP.
15
Figure 1.2 Streamlined representation of the order Chlamydiales. Each family is
represented by a different color. Organisms not colored have not yet been classified into a
family (environmental Chlamydias). However organisms with the prefix “Candidatus”
have been proposed as novel species. An asterisk (*) indicates Chlamydiales identified in
previous studies.
16
1.9. CLOs identified in epitheliocystis affected fish by antigenic and
molecular methods
1.9.1. Candidatus Piscichlamydia salmonis (CPs)
This novel Chlamydia found in AS with EP in Norway and Ireland was proposed to be a
new species in 2004 (Draghi et al., 2004). Phylogenetic analysis of this new agent
demonstrated it to be distinct from previously described CLOs (Meijer and Ossewaarde
2002; Meijer et al., 2006). CPs was also identified in AC, in fresh water production
facilities in Canada and in wild BT in a river in Switzerland. Interestingly the wild BT
were also infected with Candidatus Clavochlamydia salmonicola (CCs), another novel
Chlamydiae that has been identified in AS from Europe (Draghi et al., 2010; Karseln et
al., 2008; Mitchell et al., 2010; Schmidt-Posthause et al., 2011). Studies of these agents
demonstrated that: 1) the same type of chlamydia can infect different fish species, 2) one
fish can have a mixed infection with more than one type of Chlamydiae, 3) the discovery
of CPs in distant geographic locations as well as in fresh and salt water shows how
widely spread and highly adapted CLOs are to different environmental conditions
(Draghi et al., 2004; 2010; Karseln et al., 2008; Mitchell et al., 2010; Schmidt-Posthouse
et al., 2011).
1.9.2. Candidatus Clavochlamydia salmonicola (CCs)
This Chlamydiae was found in salmonid fish from freshwater in Norway, and is thought
that it may be a third genus in the family Chlamydiaceae, or a closely related sister family
(Karlsen et al., 2008). While CPs has been shown to infect fish in both environments, it is
17
still unknown if CCs can also infect salmonids in salt water (Karlsen et al., 2008; Draghi
et al., 2010; Mitchel et al., 2010). One systematic study in wild BT concluded that
infection with CCs progressively disappeared as the fish got closer to the ocean
(Schmidt-Posthaus et al., 2011).
1.9.3. Neochlamydia spp.
In 2007 another novel CLO was identified from the gills of AC. This new EP organism
was an uncultured Neochlamydia spp., which had 16S rDNA sequence identity to a
Neochlamydia sp. that had been obtained before in Europe from a cat with ocular disease
(von Bomhard et al., 2003; Draghi et al., 2007). This may be the first case where a
clinically relevant member of Chlamydiales has been identified from both a mammalian
and piscine source (Draghi et al., 2007). While further investigation into this link is
necessary, it may be interesting to explore if these organisms are able to cause disease in
a broad spectrum of hosts, since it has been speculated that some CLOs may be zoonotic
pathogens (Polkinghorne et al., 2010).
1.9.4. Candidatus Parilichlamydia carangidicola
This novel salt water CLO agent was identified in yellowtail kingfish from South
Australia. Based on 16S rDNA sequencing, it was proposed to be a member of a novel
family “Candidatus Parilichlamydiaceae” within the order Chlamydiales (Stride et al.,
2013).
18
1.9.5. Candidatus Actinochlamydia clariae
This organism was recently identified in juvenile African sharptooth catfish (Clarias
gariepinus) from Uganda associated with mortality events due to gill disease (Steigen et
al., 2013). The bacterium is 17.6-18.0% different from the 16S rDNA in Chlamydiaceae,
qualifying this organism as a member of the Chlamydiales, but not of the family
Chlamydiaceae. Actinochlamydiaceae currently has one genus Actinochlamydia, which
forms a sister taxon to Piscichlamydiaceae and Chlamydiaceae, and it has been proposed
to be a new family within the order Chlamydiales.
1.9.6.
Candidatus Renichlamydia lutjani
This CLO was identified in the spleen and kidney of blue striped snapper (Lutjanus
kasmira) in tropical marine water from Hawaii. This is the first, and so far only report, of
chlamydial infection affecting internal organs in fish. This agent was identified using
PCR and LM, thus more studies are needed to localize the sequences in the EP lesions
such as ISH.
1.10. Non-chlamydial agents associated with epitheliocystis
1.10.1. Candidatus Brachiomona cysticola (CBc)
This novel organism belongs to the class ß-proteobacteria, order Burkholderiales, and it
was the only agent associated with branchial intracellular microcolonies during an EP
outbreak in Norwegian AS in salt water. It was identified by PCR using universal 16S
rDNA bacterial primers and localized to inclusions by fluorescent in situ hybridization
(FISH). This study demonstrated that CLOs are not the only group of bacteria causing EP
19
(Toenshoff et al., 2012). CPs was also previously associated with EP outbreaks in this
same region (Draghi et al., 2004) and this discovery raises some interesting questions.
For example, could there be more than one organism involved in the pathogenesis of EP,
each perhaps more common at different times? Very recently, an epidemiological study
using qPCR found that CBc is a common agent of EP in seawater-farmed AS in Norway
and Ireland but that CPs were also identified at relatively low levels of infection,
apparently independent of EP prevalence (Mitchell et al., 2013). The morphology of CBc,
based on EM, was briefly described to be pleomorphic with a cell wall typical of a Gramnegative bacterium. There is presently no evidence available for its transmission, life
cycle or pathogenesis since the organism has not been cultured.
1.10.2. Endozoicomonas elysicola (Ee)
This Gram-negative rod, a strictly aerobic γ-proteobacteria, was isolated from the ornate
leaf slug (Elysia ornata) from Japan on marine agar at 25 °C (Kurahashi and Yokota,
2007). More recently this bacterium was found localised to branchial EPI and were
considered to be the cause of EP in cobia larvae from Colombia (Mendoza et al., 2013).
1.10.3. Rickettsia-like organisms (RLOs)
Rickettsiales belong to the α-proteobacteria class and generally survive only as
endosymbionts of other cells (Garrity, 2011). Evidence associating RLOs in fish with EP
is based solely on EM studies. Due to the pleomorphic shape of the agent, e.g. EP in
American plaice, the ultrastructure was considered to be closer to RLO than CLO
20
(Morrison and Shum, 1983). There is no molecular or immunological evidence that links
these bacteria to inclusions seen in EP.
1.10.4. Other agents
Viruses, such as a herpesvirus, a paramyxovirus, an iridovirus,
bacteria, such as
flexibacter-like bacteria, Aeromonas salmonicida, Vibrio alginolyticus, Vibrio vulfnificus,
the myxozoan Tetracapsuloides bryosalmonae, protozoa such as Trichodina spp., and
metazoans, such as Gyrodactylus spp., Microcotyle spp., and Lamellodiscus spp., have all
been identified to co-infect fish affected with EP. However, their role in the pathogenesis
of the observed gill lesions and population morbidity and mortality events is unknown
(Hoffman et al., 1969; Wolke et al., 1970; Paperna, 1977; Paperna et al.,1978; Paperna
and Baudin, 1979; Zachary and Paperna, 1977; Miyazaki et al., 1986; Bradley et al.,
1988; Bradley et al., 1989; Turnbull et al., 1989; Anderson, 1992; Padros and Crespo,
1995; Nylund et al., 1998; Szakolczai et al., 1999; Kvellestad et al., 2005; Meijer et al.,
2006 ; Steinum et al., 2009; Schmidt-posthaus et al., 2011, Toenshoff et al., 2012).
Several authors have noticed a temporal correlation between the presence of
epitheliocystis inclusions (EPI) and monogenean infestations, and these investigators
have postulated a link between the two infections (Anderson, 1992; Padros and Crespo,
1995; Nylund et al., 1998; Kvellestad et al., 2005), although both infections would be
considered common in some wild and captive fish populations.
21
1.11. Transmission and pathogenesisIt is possible that the EP agent, especially if
it is an obligate intracellular organism, could infect other vertebrate, invertebrate or
protistan hosts that may act as reservoirs. It has been found that Parachlamydia spp. and
Neochlamydia spp. can infect and replicate within protists such as Acanthamoeba spp.
and Hartmannella spp. suggesting that these fauna may serve as a reservoir for some
agents that have been associated with EP in fish (Amann et al., 1997; Draghi et al., 2004;
Collingro et al., 2005; Corsaro and Greub, 2006). However, there has been only limited
investigation to date regarding the identification of associated agents outside the fish.
DNA sequences consistent with CLOs were not detected from water samples taken from
facilities with fish affected by EP, suggesting either that the agent was not in the water
column at the time or the water column and its associated protistan fauna may not serve
as a reservoir of CLOs (Draghi et al., 2010).
On the other hand, studies conducted with wild BT in Swiss rivers and a survey of
salmonid pathogens in ocean-caught fish in British Columbia, Canada, suggested that
wild fish populations may serve as an alternate environmental reservoir host for EP
infections that occur on fish farms (Kent et al., 1998; Schmidt-Posthaus et al., 2011). The
Swiss rivers, from which wild BT were diagnosed with EP, had no aquaculture facilities,
indicating that wild fish populations can be naturally infected without the presence of or
interaction with farmed fish (Schmidt-Posthaus et al., 2011). Additionally, no EP
outbreaks in the waters off the Canadian Pacific northwest coast had been reported in
22
pen-reared salmonids, and yet, 29 wild marine fish species were found to have EPI
suggesting that wild fish may serve as a reservoir of EP (Kent et al., 1998).
Successful isolation and culture of Chlamydiae or other possible EP agents, such as
RLOs or Burkolderiales from infected fish, would allow the establishment of
experimental models to test if EP bacteria are limited to fish or if they are capable of
utilizing amoebas or other organisms in the environment as reservoirs (Draghi et al.,
2010).
In general, there is insufficient information regarding the developmental cycle and
pathogenesis of most of the organisms of Order Chlamydiales. However, for the genus
Chlamydia, which contains well-known human pathogens such as Chlamydia
trachomatis, their pathogenesis is better understood, in part as cell culture methods have
been developed. Novel host-pathogen interactions have been revealed by studying how
C. trachomatis virulence factors manipulate eukaryotic cellular function and how these
factors are translocated into the host cell (Kenneth and Hackstadt, 2002; Kumar and
Valdivia, 2008; Cocchairo et al., 2008; Jorgensen and Valdivia, 2008; Saka and Valdivia,
2011; Scidmore, 2011).
As obligate intracellular parasites, Chlamydiae have evolved unique adaptations for
intracellular parasitism. It has been demonstrated that Chlamydia trachomatis resides
within a membrane-bound compartment or ‘inclusion,’ and from this location the
23
pathogen manipulates the actin cytoskeleton and microtubule-based motors, inhibits
lysosomal recognition of the inclusion, activates signaling pathways, re-routes lipid
transport, and prevents the onset of programmed cell death (Kumar and Valdivia, 2008;
Cocchairo et al., 2008; Jorgensen and Valdivia, 2008; Saka and Valdivia, 2011;
Scidmore, 2011). Additional studies to prove that CLOs cause the same effects in fish
host cells and to understand the impact that novel CLOs derived from fish could have in
other animals, including humans, are needed, but have been limited due to our collective
inability to isolate and maintain CLOs associated with EP under in vitro conditions
(Draghi et al., 2007).
The developmental cycle of Chlamydiae is another important adaptation to survive the
hostile extracellular environment and overcome intracellular mechanisms that would
prevent parasitism. The cycle may be considered to be superficially analogous to
bacterial sporulation since it has an environmentally stable cell type, the elementary body
(EB), and a functionally and morphologically distinct vegetative cell type called the
reticulate body (RB) (Beatty et al., 1994; AbdelRahman and Belland, 2005). The primary
function of the EB is to survive in the extracellular environment for enough time to
invade a susceptible host cell, while RBs are the metabolically active and replicating
stage (Figure 1.2).
In the sea bream, two types of developmental cycles have been described (Crespo et al.,
1999). Cycle I, is similar to the typical Chlamydiae developmental cycle including EB,
24
RB and intermediate body (IB) stages. Cycle II is dissimilar and possesses other types of
stages described as primary long cells, intermediate long cells, non-vacuolated small cells
and vacuolated small cells (Crespo et al., 1999). Cycle II exhibits ultrastructural
similarities to Rickettsiae with a CLO-like developmental cycle, e.g. Ehrlichia
ruminantium (Crespo et al., 1999).
There were ultrastructural differences noted between developmental stages present in the
inclusions of Norwegian AS with proliferative gill lesions and Irish AS with no
proliferative lesions (Draghi et al., 2004). While the Norwegian fish presenting with
proliferative lesions had IBs and RBs within the inclusions, the Irish fish without
proliferative lesions had EBs, RBs, IBs and another cell type, described previously only
in salmonids as ‘head-and-tail forms’ (Rourke et al., 1984; Bradley et al., 1988). This
may suggest that the two different CLO developmental cycles are representative of two
different bacterial species affecting farmed AS; one occurring in proliferative gill lesions
and one occurring in non-proliferative lesions (Crespo et al., 1999; Draghi et al., 2004).
However, the Chlamydia trachomatis model has suggested that distinct stages of
chlamydia development are undoubtedly triggered in response to changes in the
intracellular environment as the infection progress (Beatty et al., 1994; Nguyen et al.,
2011), and this could also explain some of the differences in CLO cycles in fish.
Finally, ultrastructural studies on AS samples containing the ß-protobacterium CBc
demonstrated large membrane-bound cytoplasmic inclusions containing pleomorphic
bacterial cells. Rounded to elongated forms approximately 0.2-0.4 µm in size and some
25
small vesicles within rounded cells where observed. Most of these morphotypes
resembled IBs while a few resembled RB forms of the CLO developmental cycle;
however, EB-like morphotypes where not observed (Toenshoff et al., 2012).
1.9. Control and treatment
Since more than one organism might be involved in an outbreak and since diagnosis can
be problematic, there is no established treatment for EP. Alternative husbandry methods,
such as decreasing stress factors and increasing water quality, around the time of
outbreaks times has been recommended (Groff, et al., 1996). Ultraviolet irradiation of
water supplies was reported to control EP outbreaks in amberjack and coral grouper
(Cephalopholis miniata)(Miyaki et al., 1998).
Assuming that the causative agents of EP are CLO and knowing that chlamydiae in
general are sensitive to antibiotics that inhibit protein production, such as tetracyclines
and macrolides, antimicrobial therapy has been attempted. Successful response to oral
oxytretracycline in largemouth bass (Micropterus salmoides) affected by EP has been
reported (Goodwin et al., 2005). No other examples of successful antimicrobial treatment
have been published. Enrofloxacin has been used to treat Chlamydia in humans; however,
this and many other antimicrobials are prohibited in fish farmed for human consumption
where the highest production losses are observed (Nowak and LaPatra, 2006),
26
1.13. Epitheliocystis in lake trout
In salmonids, EP has been reported in AS, AC, rainbow trout (Oncorhynchus mykiss), LT
and BT, Chinook (Oncorhynchus tshawytscha), chum (Oncorhynchus keta), coho
(Oncorhynchus kisutch), pink (Oncorhynchus gorbuscha) and sockeye salmon
(Oncorhynchus nerka) (Rourke et al., 1984; Bradley et al., 1988; Kent et al., 1998;
Nylund et al., 1998; Gavruseva, 2009; Schmidt-Posthaus et al., 2012; Toenshoff et al.,
2012) principally in Europe and North America (Tables 1.1 and 1.2). Only in AS, AC and
BT, have agents associated with EP been identified using molecular methods; CPs in AS,
AC and BT; CCs in AS and BT; and CBc in AS. For the seven other salmonid species,
only the lesions and morphological characteristics of the causative agent have been
described using LM and EM (Rourke et al., 1984; Bradley et al., 1988; Nylund, 1998;
Kent et al., 1998).
In LT, EP has been reported from two epizootics in yearling juveniles that occurred in
hatcheries in the Great Lake region (McAllister and Herman, 1989; Bradley et al., 1988).
While EP was present during these epizootics, the causative agent of the mortality events
was later considered to be a herpes-like virus and the condition was termed epizootic
epitheliotropic disease (EED) (Bradley et al., 1989). This conclusion was based on
experimental studies with EED virus (EEDV), in which horizontal transmission from
infected fish or with skin scrapings/homogenates was demonstrated (Bradley et al., 1989;
McAllister and Herman, 1989; Kurobe et al., 2009).
27
Table 1.1 Epitheliocystis reported in farmed salmonids.
Species
Reports
Location
Water
Atlantic salmon
5
Norway
Norway & Ireland
Norway
Norway
Norway
Seawater
Seawater
Seawater
Fresh
Sea water
References
Nylund et Kvenseth,
1998
Draghi et al., 2004
Karlsen et al., 2008
Mitchell et al., 2010
Toenshoff et al., 2012
USA & Canada
USA & Canada
Fresh
Fresh
Draghi et al., 2007
Draghi et al., 2010
USA
Fresh
Rourke et al., 1984
Fresh
Bradley et al., 1988
2
Artic charr
Rainbow trout
1
1
Lake trout
USA
Table 1.2: Epitheliocystis reported in wild salmonids.
Species
Reports
Location
Water
References
Chinook salmon
1
Canada
Seawater
Kent et al., 1998
Chum salmon
1
Canada
Seawater
Kent et al., 1998
Coho salmon
1
Canada
Seawater
Kent et al., 1998
Pink salmon
1
Canada
Seawater
Kent et al., 1998
2
Norway
Switzerland
Fresh
Fresh
Karlsen et al., 2008
Schmidt-Posthaus et al.,
2011
1
Russia
Fresh
Gavruseva, 2009
Brown trout
Sockeye salmon
28
1.14. Conclusion
Multiple organisms belonging to the Order Chlamydiales have been found to be putative
causative agents of EP in numerous fish species, including salmonids. However, there has
been recent evidence that other infectious agents such as CBc and Ee also are associated
with EP outbreaks, challenging the theory that CLOs are the sole etiological agents of EP
(Toenshoff et al., 2012, Mendoza et al., 2013). Furthermore, the multitude of species that
have been related to EP might explain the reported variation in the prevalence, host
species, geographic distribution, and disease severity (Frances et al., 1997). Additional
research focused on the identification of the microorganisms involved and their means of
culture is needed in order to finally define which agents are actually causal in the
pathogenesis of EP and are associated with clinical signs and mortalities.
1.15. Rationale
Lake trout are one of the 18 species of fish belonging to the Salmonidae family that are
found in Ontario. They are present in the Great Lakes and other lakes on the Canadian
Shield with cold, deep and well-oxygenated waters (OMNR, 1993). Lake trout are
important for recreation and food, since it is one of the highest quality angling fish in the
province. They are also used as a biological indicator of a healthy aquatic environment
due to their high sensitivity to environmental changes in comparison with other fishes
(OMNR, 1993).
The Ontario Ministry of Natural resources (OMNR) operates nine provincial fish culture
facilities, producing approximately 8 million fish each year. In 2011, 3.3 million of these
29
were LT, the majority of which were stocked into the Great Lakes. This effort intends to
restore populations that were destroyed by sea lamprey predation and overfishing in the
1950s (OMNR, 2010). A substantial portion of LT grown by the OMNR are raised at the
Blue Jay Creek (BJC) Fish Culture Station. The staff there, in collaboration with the Fish
Pathology Laboratory (FPL) at the University of Guelph, have identified that the yearly
mortalities experienced during EP outbreaks are a serious health problem.
1.15.1. Purpose
To identify the infectious agents associated with a case of EP using PCR and laser
capture microdissection (LCM). To describe the lesions associated with EP using LM and
TEM, and determine the lesions associated with the clinical presentation and mortality.
1.15.2. Hypothesis and objectives
Despite the detection of EP in more than 50 fresh water and marine fish species, the
presence of CLO using molecular techniques (PCR/ISH/LCM) has been demonstrated in
only seven of these fish hosts: AS, AC, barramundi, silver perch, BT, leopard shark,
yellowtail kingfish and cobia. Since these studies have shown that bacteria from the order
Chlamydiales are a cause of EP, it washypothesized that:
i)The causative agent(s) of epitheliocystis outbreaks in lake trout are members of the
order Chlamydiales and ii) The Chlamydiales found in clinically affected lake trout are
highly similar (90 % nucleotide identity) to either CPs or CCs.
30
Polymerase chain reaction, TEM, bacteria culture, and LM, were therefore used to
establish the presence of detectable organisms in gill tissue from LT with EP. The first
objective was to identify potential EP agents using universal bacterial and Chlamydialesspecific primers that amplify regions of the 16S rRNA gene (Weisburg et al, 1991;
Relman, 1993; Greisen et al., 1994; Fukatsu & Nikoh, 1998; Everett et al., 1999;
Ossewaarde and Meijer 1999). The Fish Pathology Laboratory has used these primers on
infected gills of AC and koi (unpublished), successfully amplifying Neochlamydia sp.
and CPs, respectively. PCR on bacterial DNA obtained from fresh gill tissue, bacterial
DNA obtained from inclusions using LCM and from bacteria cultured from affected gills
will be performed using the primers previously mentioned (Weisburg et al, 1991;
Relman, 1993; Greisen et al., 1994; Fukatsu & Nikoh, 1998; Everett et al., 1999;
Ossewaarde and Meijer 1999). Bands generated will be sequenced in order to determine
if the EP agent in LT is genetically related to other known Chlamydiales such as CPs and
CCs known to infect fish. Gills from EP+ cases were used for the second objective,
which was to localise the organisms identified to EPI using TEM, immunohistochemistry
and LCM.
Even though EP has been associated with mortalities at BJC, there is little evidence to
link the presence of EP to clinical signs in fish. No agent has yet been cultured despite
numerous efforts, and consequently Koch postulates have not been fulfilled. In an attempt
31
to associate the presence of EPI and mortality in LT BJC population a second hypothesis
was therefore proposed:
Peak mortality in lake trout is positively associated with branchial lesion severity and the
number of EPI
The first objective for this second hypothesis was to identify cases of gill disease in LT
with EPI. Light microscopic lesions seem to be fairly characteristic in fish examined at
the FPL; however, the inclusions are not consistently present. Tissues of affected fish
from 2006 to 2013 were archived in formalin and less commonly were also frozen. The
second objective was to collect gill samples at different time points during a single
outbreak from which mortality data was available. These resources will be used to
systematically describe lesion severity and the number of colonies in affected fish over
time.
32
2.
MATERIALS AND METHODS
2.1. Background
Lake trout were obtained from the BJC Fish Culture Station. The hatchery has a singlepass, flow-through system with ground water and multiple 6000 L indoor tanks. There
are about 6200 yearlings contained in each tank, held at ~ 8 ºC and exposed to a natural
day/night light cycle. When an EP outbreak is suspected, the staff rules out parasites and
BGD using toluidine blue-stained wet mounts. If hyperplasia and necrosis are observed
and no agents are identified, fish are sent to the Fish Health Laboratory (FHL) at the
Science complex (SCIE) laboratory 3207, University of Guelph. Fish are tested using
standard bacteriology and virology methods in order to obtain an etiologic diagnosis.
When an etiologic diagnosis is not made, and histology diagnosis is required, the FHL
will send samples to the FPL, where a diagnosis of EP may be made using LM based on
the presence of gill lesions.
2.2. Retrospective, prospective sample collection and tissue processing
The majority of fish tissues from affected lake trout were archived in formalin between
2006 and 2009. Since 2010, live fish with clinical histories and signs suspicious for EP
were submitted for diagnosis directly to the FPL (Table 2.1). In 2013 four groups of fish
were collected from Raceway 8 (R8): Group A consisted of five fish from the preclinical
period collected on December 3rd; Group B consisted of five fish from the early clinical
period collected on January 3rd Group C, consisted of ten fish from the clinical period
33
collected on January 8th and finally Group D had ten fish collected during peak mortality
on January 18th (Group D). A total of 30 fish, therefore, were collected for histological
and other evaluations as outlined below. Fish from R8 were selected as they had not been
treated during the disease outbreak. Fish were euthanized at the FPL by an overdose of
benzocaine (Aqualife TMS, Syndel Laboratories Ltd, Qualicum Beach, British Columbia,
V9K 1V5, Canada), and dead fish were immediately placed on ice.
The 2013 samples were processed as follows (Appendices 7.1: EP-PM protocol1). Before
opening the carcass, three pieces of dorsal skin and dermis were excised and either fixed
in 10 % buffered formalin (ACP Chemicals Inc., Saint-Léonard, Québec, H1R 1A5,
Canada), frozen at -80 ºC or placed in RNAlater. The first (outermost) gill arch was
bisected; one half was placed into RNAlater in a 1.5 ml microcentrifuge tube and the
other half was placed in formalin. Half of the second arch was frozen at -80 ºC and the
other half was formalin-fixed, the third arch was placed in RNA later (cut in three pieces)
and finally the fourth arch was frozen. The same procedure was then applied to the
opposite gill arches. Heart, liver, spleen, head and tail kidney were frozen at -80 ºC and
fixed in formalin to allow identification of other diseases. Between fish the dissections
tools were immersed in 70 % ethanol and flamed to prevent cross contamination. Before
2013, if freshly dead fish were available, two gill arches were excised and stored at -80
ºC, the coelomic cavity was opened and the whole carcass was immersed in 10 % neutralbuffered formalin for at least 24 h.
34
Table 2.1 Lake trout epitheliocystis cases submitted to the Fish Pathology
Laboratory between 2007 and 2013.
Case
number
B126-07
Date
Tank/lot
Number
March-April, 2006
S6-S7
72 in formalin
B037-10
February 24, 2010
342/150
6 live, 6 in formalin =12 (fry)
B047-10
March 25, 2010
342/150
8 live, 6 in formalin =14 (fry)
B028-11
February 11, 2011
R1
3 fish in formalin (fingerling)
B034-11
Feb 18, 2011
R1
6 live (fingerling )
B042-11
March 3, 2011
R1
8 live (fingerling )
B214-11
December 21, 2011
R4
6 live (fingerling )
B016-12
January, 2012
R4
10 frozen fish and 10 formalin fixed
gills (fingerling )
B016-12
February, 2012
R4
10 frozen fish and 10 formalin fixed
gills (fingerling )
B014-13 (A)
December 3, 2012
R8
5 frozen fish and 10 formalin fixed
gills (fingerling )
B002-13 (B)
January 3, 2013
R8
5 frozen fish and 10 formalin fixed
gills (fingerling )
B002-13 (C)
January 8, 2013
R8
10 live (fingerling )
B015-13 (D)
January 18, 2013
R8
10 frozen fish and 10 formalin fixed
gills (fingerling )
35
2.3. Bacterial isolation
To examine the possibility that the EP agent is a facultative intracellular bacterium,
BFCG plates, previously used to isolate Piscirickettsia salmonis (Mauel et al., 2008) was
trialed for group C (ten fish) some of the 2013 samples. The BFCG media contained 3 %
fetal bovine serum, 0.1 % cysteine, 1 % glucose in 5 % sheep blood. Prior to DNA
extraction, the gills were homogenized using a RNA/DNase-free disposable sterile pellet
pestle 1.5µl (VWR, 2360 Argentina Road, Mississauga, ON, Canada). The homogenized
tissue was streaked onto BFCG, BA (blood agar) and TSA (Trypticase soy agar) plates
and incubated at 10 and 21 ºC.
2.4. Histopathology
Formalin-fixed gills and organs from larger fish were trimmed into histology cassettes.
Fingerlings were dissected transversely in segments of approximately 4 mm, and fry were
cut longitudinally in two halves using a scalpel blade. Gills arches were dissected and
placed in the cassette separate from the body. Tissues were then submerged in Surgipath
Decalcifier II solution (Surgipath Medical Industries Inc., Richmond, Illinois, 600717702, USA) to decalcify for about 2 h. Afterwards segments were processed routinely for
paraffin embedding, sectioned at 4 μm, and sections were mounted on glass slides and
stained with hematoxylin and eosin (H&E) (Animal Health Laboratory, Guelph, Ontario,
N1G 2W1, Canada). Pierce Van der Kamp (PVK), Wolbach Giemsa and B&H (Brown
and Hopps) were used selectively (see results).
36
2.4.1. Scoring of gill morphology
The common pathological lesions observed in selected LT EP cases from 2013 were
semi-quantitatively scored. Lesions scored included: lamellar epithelial hyperplasia (LH),
goblet cell metaplasia (GM), the number of eosinophilic granular cells (EGC), cell
necrosis (N) and the presence of intracellular bacterial inclusions (EPI). The criteria and
scoring were established first by comparing between less affected fish from December 3rd
(Group A) and fish with the most marked lesions from January 8th (Group C), and then all
groups were reassessed (Table 2.2). The common pathological lesions were assessed in
five fish (five gill arches) per group. LH was scored in five filaments per arch at 4X and
in the case of EGC, GM, N and EPI, a total of ten fields at 40X were evaluated. A portion
of the gills were then scored by a separate person. Differences in lesion scores were
examined by one-way ANOVA, and statistically significant differences were determined
using Tukey’s HSD. The software utilized was SAS/STAT (version 9.2, SAS Institute
Inc., Cary, NC, USA).
37
Table 2.2 Semi-quantitative scores in lake trout gill lesions using light microscopy (modified from Mitchell et al.,
2012).
Pathology
(Score)
None
(1)
Mild
(2)
Moderate
(3)
Severe
(4)
Lamellar
hyperplasia (LH),
4X*
Necrotic cells
(N), 40X*
Epitheliocystis
Metaplastic goblet
inclusions (EPI), 40X* cells (GM), 40X*
Eosinophilic granular
cells (EGC), 40X*
Not evident or very
minor
0-1
0
0
0
<10 % of the filament
view affected
2-5
1-3
1-4
1-4
10 %-50 % of the
filament affected
6-10
4-9
5-10
5-10
>50 % of the filament
affected
>10
>10
>10
>10
* Magnification used to assess each lesion
38
2.5. DNA extraction
2.5.1. Gill tissue
DNA extraction from frozen gill tissue preserved at -80 ºC was performed following the
manufacturer’s (Qiagen DNeasy) mouse-tail protocol. The genomic DNA obtained was
stored at -20 ºC until use. Positive and negative controls for genomic DNA extraction
were gills previously determined to be PCR-positive for CPs and liver tissue from healthy
rainbow trout PCR-negative for bacterial DNA, respectively. Liver was selected as a LT
genomic DNA negative control due to the low risk of contamination with prokaryotic
bacterial DNA.
2.5.2. Bacterial isolates from gill tissue
DNA extraction was performed based on the InstaGene™ matrix protocol for bacteria
(Bio-Rad Laboratories, Hercules, California, 94547, USA). Briefly, pure colonies were
suspended in 1 ml of sterile Milli Q water in a 1.5 ml microcentrifuge tube. The tubes
were centrifuged for 1 min at 12,000 rpm (Centrifuge 5424, Eppendorf NA, Inc.,
Westbury, New York 11590, USA), followed by the removal of the supernatant. Then
200 μl of InstaGene™ was added to the residual pellet in each tube and incubated in a
heat block at 56 °C for 20 min with shaking at 600 rpm (Thermomixer, Eppendorf,
Hauppauge, New York, 11788, USA). After incubation, tubes containing the samples
were vortexed at high speed for 10 sec and placed in a heat block at 99 °C with shaking at
600 rpm for 8 min. Then tubes were vortexed again for 10 sec and centrifuged at 12,000
rpm for 3 min. Approximately 180 μl of supernatant was passed to a new microcentrifuge
tube and stored at -20 °C until needed. Vagococcus salmoninarum isolated from Atlantic
39
salmon (FPL B016-10) was used as positive controls for DNA extraction and the PCRs.
An empty microcentrifuge tube was also processed as a negative control.
2.5.3. DNA extraction from formalin-fixed paraffin-embedded gill tissues
Genomic DNA was extracted from gill samples with EP lesions using LCM. A 7 μm
thick section was cut from each paraffin block of interest, and the section was mounted
on glass slides (Fisherbrand®, Fisher Scientific, Fair Lawn, New Jersey, 07410, USA)
without baking or staining. LCM was performed to retrieve cells containing EPI using an
inverted microscope system and a 15 μm diameter laser (Olympus IX50/IX70). Staining
of the tissues was achieved by immersion in xylene (Sigma-Aldrich, St. Louis, MO,
USA) for 5 min (two times), 100 %, 95 % and 70 % ethanol washes, DEPI
(diethylpyrocarbonate) water wash, hematoxylin (Fisher®) for 2 min, DEPI water wash,
ammonia water (bluing reagent) for 60 s, 70 % and 95 % ethanol washes, eosin (Fisher®)
for 2 min, 95 %-100 % ethanol wash and a final xylene immersion for 1 min. The resin
layer containing tissues was removed from HS LCM caps (Arcturus® Cap Sure®) using
sterile forceps and placed in 1.5 ml DNAse/RNAse-free microcentrifuge tubes. DNA was
extracted from the resulting pellet using a DNA extraction kit for formalin-fixed paraffinembedded (FFPE) tissues following the suggested protocol (QIAamp® DNA FFPE
Tissue, Qiagen GmbH, D-40724 Hilden). Briefly, tissue was lysed with 180 μl Qiagen
ATL tissue lysis buffer and 20 μl proteinase K, vortexed and incubated at 56 °C for 24 h.
Samples were heated to 90 °C for 1 h, then centrifuged at 6,000 g for 2 min. Extraction
was conducted as per the manufacturer’s directions; the DNA was diluted in AE buffer
(Qiagen) and stored in 1.5 ml microfuge tubes at –20 °C until used.
40
Finally. for all nucleic acid extraction protocols, the DNA concentration was quantified
using a Nanodrop (NanoDrop™ 2000, Thermo Scientific). The purity of the DNA was
assessed by the 260/280 and the 260/230 ratios, which should be close to 1.8 and between
2.0-2.2, respectively.
2.6. Polymerase chain reaction (PCR)
2.6.1. Primers
Two different primer pairs were used in this study (Table 2.3), i.e. the first to amplify 16S
rDNA from conserved regions common to Order Chlamydiales and the second to amplify
16S rDNA regions common to all bacteria. Primers for all PCR experiments were
synthesized by the Guelph Molecular Supercenter (Laboratory Services Division,
Guelph, Ontario, Canada) and diluted to a stock concentration of 100 pmol/l.
Chlamydiales primers were tested first on their ability to amplify ~300 bp, ~800 bp and
~1500 bp gene fragments from positive controls such as CPs and Chlamydophila abortus.
Alternatively, a Chlamydiales-specific primer set that produced a 270 bp product, FOR2
and REV2 (Ossewaarde and Meijer, 1999), were used on samples when the first set was
not able to produce amplicons (Figure 2.1).
Universal bacterial primers targeting the 16S rDNA went through the same validation
processes to which Chlamydiales primers were subjected. Primers were tested first on
their ability to amplify ~300 bp, ~800 bp and ~1500 bp gene fragments from the positive
control Vagoccoccus salmoninarum. These primer sets were used to produce ~370 bp,
41
~800 bp and ~1500 bp amplicons of genomic DNA obtained from FFPE tissue, fresh gill
tissue and isolated bacterial colonies, respectively (Table 2.3).
EP in LT was first reported by Bradley et al. (1988) from fish also affected by EEDV.
Therefore, LT from BJC were tested for the presence of this virus using PCR (Kurobe et
al., 2009) (Table 2.3). DNA from tissue known to be infected with KHV (another fish
herpes virus) was the positive control for the PCR.
2.6.2. Amplification protocols
All PCR products were amplified in individual 50 μl reaction mixtures containing 1000
ng of DNA, 25 μl of 2x TopTaq Master Mix (contains TopTaq DNA Polymerase,
TopTaq PCR Buffer with 3 mM MgCl2 and 400 μM each dNTP) (Qiagen), 2 ul of each
forward and reverse primer (diluted to 20 pmol/μl working solution), 5 μl of 10x Coral
Load Concentrate (Qiagen) and 9 μl of RT-PCR grade water (Ambion), using thermal
cyclers (Mastercycler personal, Eppendorf). Location of ~300 bp 16S rDNA products.
The 16SIGF and 16SIGR primer pair was used in a touchdown PCR (Appendix 7.3:
protocol CLORX1) to limit secondary priming. Reactions were begun by a 15-min
incubation at 94°C, followed by 40 cycles, each consisting of denaturation at 94 °C for 45
s, primer annealing for 45 s, and extension at 72 °C for 45 s. Annealing temperatures
were initiated at 66 °C and decreased by 1 °C every third cycle until 61 °C, at which
temperature the final 25 cycles were performed. After 40 cycles, a 7-min extension step
at 72 °C was performed. To obtain ~800 bp 16S sequence products, PCR reactions using
16SIGF and 806R primer pairs set (Appendix 7.3: protocol CLORX2) were initiated with
42
a 15-min incubation at 94 °C, followed by 40 cycles, each consisting of denaturation at
94 °C for 30 s, annealing at 55 °C for 45 s, and extension at 72 °C for 45 s, with a final
extension of 7 min. For near-full-length 16S rRNA amplicons (~1500 bp) produced by
the 16SIGF and 16SB1 primer pair, the PCR protocol reaction (Appendix 7.3: protocol
CLORX3) was initiated by incubation at 94 °C for 15 min, followed by 40 cycles, each
consisting of denaturation at 94 °C for 40 s, annealing at 58 °C for 40 s, and extension at
72 °C for 7 min. To amplify 16S rRNA genes with universal primers, PCR conditions
were as follows: 5 min incubation at 95 °C, followed by 35 cycles of 30 s at 94 °C, 60 s
at 56 °C, and 60 s at 72 °C. A 10 min final step at 72 °C was performed to extend any
incomplete amplicons. Two types of negative controls were run in the PCR reactions, i.e
the negative DNA extraction and negative PCR template controls consisting of 45 μl
from the mastermix plus 5 μl RT-PCR grade water (Ambion). A touchdown PCR
protocol was utilized for REV2 and FOR2 primers (Appendix 7.3: protocol CLORX4)
which consisted of 10 min incubation at 95 °C and denaturation at 95 °C for 60 s.
Annealing temperatures began at 62 °C for 60 s and decreased by 1 °C every two cycles
until 58 °C, at which temperature the final 40 cycles were performed. Finally a 7-min
extension step at 72 °C was required.
In the case of the 16S universal bacterial primers, two PCR protocols UNIRX1 and
UNIRX3 (Appendix 7.3), were used to amplify the ~370 and ~1500 bp products
respectively. These two protocols have the same steps, which consisted of a 5-min initial
denaturation at 95 ºC, followed by 30 cycles each consisting of denaturation at 96 ºC for
30 s, annealing at 60 ºC for 60 s, extension at 72 ºC for 60 s. A final extension of 10 min
43
at 72 ºC was performed. The protocol to amplify a ~800 bp product (Appendix 7.3:
protocol UNIRX2) initiates with incubation at 94 ºC for 5 min, followed by 30 cycles of
denaturation at 96 ºC for 15 s, annealing at 60 ºC for 1.5 min, extension at 72 ºC for 2
min. A 5 min final step at 72 °C was performed.
Screening for EEDV from DNA extracted from gill samples infected with EP, using 339F
and 340R primers was also performed (Appendix 7.3: EEDVRX1 protocol). Initial
denaturation was at 95 ºC for 5 min, followed by 30 cycles of denaturation at 95 ºC for 30
s, with annealing at 60 ºC for 30 s, and finally extension at 72 ºC for 30 s. When the
cycles were done, a final extension at 72 ºC for 5 min was performed.
44
Figure 2.1 Chlamydiales primer localization along the 16S rRNA gene of positive
control Candidatus Piscichlamydia salmonis.
45
Table 2.3 List of primers used in this study.
Product size
Primer
16SIGF
Chlamydiales 16rRNA
signature (298 bp)
Chlamydiales 16rRNA
signature (270bp)
Sequence
5'-CGGCGTGGATGAGGCAT-3'
Position
References
40-57
Everett et al., 1999
16SIGR
5'-TCAGTCCCAGTGTTGGC-3'
FOR2
5'-CGTGGATGAGGCATGCAAGTCGA-3'
REV2
5'-CAATCTCTCAATCCGCCTAGACGTCTTAG-3'
16SIGF
5'-CGGCGTGGATGAGGCAT-3'
340-323
~1-10
Ossewaarde
& Meijer, 1999
~280-270
40-57
Everett et al., 1999
806-787
Relman, 1993
40-57
Everett et al., 1999
1527-1505
Fukatsu & Nikoh,
1998
Chlamydiales 16 Sr RNA
half-length (766 bp)
806R
Chlamydiales 16Sr RNA
near-full-length (1487 bp)
5'-GGACTACCAGGGTATCTAAT-3'
16SIGF
5'-CGGCGTGGATGAGGCAT-3'
16SB1
5'-TACGGYTACCTTGTTACGACTT-3'
46
Table 2.3 (Continued from page 46) List of primers used in this study.
Product size
Universal 16Sr RNA
signature (380bp)
Primer
DG74
Universal 16SrRNA
signature (~1500bp)
EEDV primers
(322bp)
5'-AGGAGGTGATCCAACCGCA-3'
Position
References
1522-1540
Greisen et al.,
1994
RW01
Universal 16Sr RNA halflength (~800bp)
Sequence
U1
5'-AACTGGAGGAAGGTGGGGAT-3'
5'-ACGCGTCGACAGAGTTTGATCCTGGCT-3'
1170-1189
1-27
Relman, 1993
U1R
5'-GGACTACCAGGGTATCTAAT-3'
806-787
27f
5'-AGAGTTTGATCMTGGCTCAG-3'
27-46
Weisburg et al,
1991
U1492R
5'-GGTTACCTTGTTACGACTT-3'
223F
5'-CATTATCGACGAAACAACG-3'
224R
5'-GTAGAAAGCCGAAACTTC G-3'
47
1492-1473
140-158
461-444
Kurobe et al.,
2009
2.7. Electrophoresis and DNA purification
Products were separated by electrophoresis in 1 % or 2 % agarose gels and visualized by
SYBR® Safe DNA gel staining (Invitrogen) and UV transillumination. Images were
recorded digitally using Gel Doc™ EZ still video image capture system (Bio-Rad
Laboratories). Only PCR reactions that amplified twice were considered for DNA
sequence analysis. All 16S rDNA PCR products were excised and purified from agarose
gels (Direct-Gel-Spin™ DNA recovery kit, LAMDA Biotech, St. Louis, MO 63011,
USA) before submission for direct sequencing (Molecular Biology Unit within
Laboratory Services of the University of Guelph).
2.8. DNA sequencing and analysis
Nucleotide sequences of amplicons were generated by oligonucleotide-directed
dideoxynucleotide chain termination DNA sequencing reactions using the respective
amplification primers (Guelph Molecular Biology Unit, Laboratory Services Division,
Guelph, Ontario, Canada). Approximately 3-20 ng of template DNA and the ABI Prism
BigDyeR Terminator Cycle Sequencing Ready Reaction kit v3.1 (Applied Biosystems,
Foster City, CA) were used for sequencing reactions. Sequencing was performed on a
GeneAmpR PCR System 9700 or 2720 Thermal Cycler (Applied Biosystems). The
BigDyeR Terminator v3.1 Cycle Sequencing Kit Protocol (Applied Biosystems) was
followed to set up and conduct the cycle sequencing reactions. Dye terminators were
removed from cycle sequencing reactions using Multiscreen-HV plates (Millipore, 75
Mississauga, ON) loaded with Sephadex G-50 Superfine (Sigma, Oakville, ON). The
48
clean reactions were electrophoresed on an Applied Biosystems 3730 DNA Analyzer
(Applied Biosystems). A minimum read length of 270 bp was generated for each of the
reactions. The chromatograms were analyzed using ABI Prism, DNA Sequencing
Analysis Software Version 3.7 (Applied Biosystems) to generate quality target sequences
within the software’s confidence range. The forward and reverse sequences were aligned
using Geneious software (Drummond et al., 2011). Standard nucleotide sequence
comparisons were made by conducting basic local alignment search tool (BLAST)
searches (Altschul et al., 1990) in GenBank to determine sequences that matched with the
highest nucleotide identity to those amplified during the investigation. Positive controls
were also sequenced and analysed in the same manner.
2.9. Immunohistochemistry
IHC was performed using the standards established by the Animal Health Laboratory
(AHL, University of Guelph, Ontario). The primary antibody was a mouse monoclonal
antibody to C. trachomatis LPS (MoAb 1631), diluted at 1:50. To see more details, refer
to Ventana ultra View™ Universal DAB Detection Kit from Roche.
2.10. Transmission electron microscopy
Various fresh tissues were fixed in cacodylate-buffered 2.5 % glutaraldehyde (pH 7.4)
and were stored at 4 °C. One filament previously processed for histopathology and
containing EPI was cut from the paraffin block and deparaffinised. Tissues were then
transferred to 1 % osmium tetroxide in cacodylate buffer and stored overnight at 4 °C.
49
The samples were dehydrated using a series of increasing ethanol concentrations (30-60
min incubation for each step; 60 %, 70 %, 80 %, 90 %, 95 %, and 100 % ethanol),
followed by 1 h in equal volumes of 100 % ethanol and propylene oxide, then 1 h in 100
% propylene oxide. Tissues were then embedded using a series of mixtures of propylene
oxide and Epon 812 (Lufts formula), e.g. 1:4, 1:3, 1:2 1:1, prepared according to the
manufacturer’s instructions (POLY BED 812 EM Embedding Media – DMP 30 Kit ,
Warrington, PA, USA) and stored overnight at 4 °C. The embedding medium was then
removed and the tissues were transferred to 100 % epon without a catalyst for 3 h. Fresh
epon with a catalyst (DMP 30, 0.4 ml/ 20 ml) was then added to the embedding capsules
for 2 h, which were then cured for 48 h at 60 °C. Thick sections (0.5 µm) were then cut
from the cured blocks with a Reichert ultra-microtome and stained with toluidine blue.
Thin sections (60 to 75 nm) from areas of interest identified by light microscopic
examination of thick sections were cut from selected areas of the block using a diamond
knife on a water surface and collected onto copper 200 mesh grids. The grids were dried
and stained with uranyl acetate and lead citrate for 10 – 30 min and examined on a
Hitachi H7100 transmission electron microscope (Hitachi High-Tech Science Systems
Inc. Ibaraki, Japan) at 75kv.
50
3. RESULTS
3.1. History and clinical findings
The BJC culture station staff has reported that gill disease outbreaks in LT are usually
associated with heavy rains during the winter. There are typically about three days when
the water becomes highly turbid and within a week the fish are affected by surface
protozoans, bacterial gill disease (BGD) and/or EP.
Clinical signs accompanying EP include the fish spreading out and staying on the bottom
of the tank/raceway, decreased feed response, reduction and loss of fright response by the
third or fourth day, lethargy, whirling and death. The typical course of the outbreak is
approximately 3 weeks, with full recovery usually in 6 weeks. Cumulative mortality can
reach ~40 %, although reduced fish growth is the principal impact and is very costly
since fish smaller than the target weight cannot be released. Besides pale gills, no other
gross lesion is notable internally or externally.
Before the 2013 outbreak, fish were moved to tank R8 on November 14th and the next
day a preventive bath treatment with formalin was applied. Samples for the present study
were taken on December 3rd, before any clinical signs were present (group A). On
December 16th there was a heavy rain event, which caused turbidity in the water for 3
days. On January 3rd the staff sent samples to the FPL to check for EP (group B), since
fish were presenting early clinical signs. The Staff also collected fish samples from R8
51
when mortality began on January 8rd (group C) and during the peak of mortality on
January 18th (group D). Total cumulative and peak daily mortality in tank R8 during the
2013 outbreak were 42.0% and 4.1%, respectively (Figure 3.1)
3.1. Bacterial isolation
A total of 16 isolates were obtained from the gills of the fish examined from 2013 (group
C) that were clinically affected with EP, the bacterial isolates were characterized
morphologically using Gram stain and identified through 16S rRNA gene sequencing
(Table 4.1). All the bacteria were Gram-negative bacilli and the predominant colonies
isolated were bacteria from the Pseudomonadaceae family. None of the isolates were
from the Burkholderiales order despite the variety of used media. Bacteria isolated on
special media BFCG at 11 ºC were Pseudomona sp. strain 8D-45 (NDB: JX905209.1)
and Serratia proteamaculans 568 strain (NDB: NR074820.1), whereas at 16 ºC, the
previously uncultured bacteria isolate, Par-w-11 (NDB: EF632915) was grown.
52
D
C
B
Days
Figure 3.1 Pattern of daily mortality of LT during winter 2013 at BJC in tank R8. The
heavy rain event occurred on December 16th (green arrow). Yellow arrows show the time
when sample were collected during the 2013 outbreaks. The first clinical signs in fish
occurred on January 3rd (B) and peak mortality on January 19th (group D). Group A
collected on December 8th not shown.
.
53
Table 3.1 Sixteen bacterial isolates derived from fish with epitheliocystis.
Isolate
Agar type &
Temperature
Accession number
Maximum
identity
Family
1
BFCG 16 ºC
EF632915: UBC, Par-w-11
99 %
Pseudomonadaceae
2
3
4
5
PS 16 ºC
PS 16 ºC
TSA 10 ºC
BA 10 ºC
EU755063.1: UBC, HM-78
100 %
Moraxellaceae
6
TSA 10 ºC
DQ068917.1: UBC, s4w18-7
99 %
Enterobacteriaceae
7
TSA 10 ºC
AY972434: Pseudomonas migulae
strain
99 %
Pseudomonadaceae
PS 10 ºC
BA 10 ºC
GU586315: Pseudomonas sp. IR-211
10
PS 10 ºC
AF320991: Pseudomonas gingeri;
11
12
TSA 10 ºC
BA 10 ºC
JX086542: UBC, LIB091_D12_1283
TSA 10 ºC
KC178584: Pseudomona cedrina
8
9
13
54
99 %
99 %
99 %
100 %
Pseudomonadaceae
Pseudomonadaceae
Enterobacteriaceae
Pseudomonadaceae
Table 3.1 (Continued from page 53) sixteen bacterial isolates derived from fish with epitheliocystis.
Isolate
Agar type &
Temperature
Accession number
Maximum
Identity
Family
14
BA 10 ºC
FJ798917.1: UBC, IC88
99 %
Aeromonadaceae
15
BFCG 10 ºC
99 %
Pseudomonadaceae
16
BFCG 10 ºC
99 %
Enterobacteriaceae
JX905209.1: Pseudomonas sp.
NR 074820.1: Serratia
proteamaculans 568 strain
UBC: Uncultured bacterial colonies; BFCG: Blood, fetal bovine serum, cysteine and glucose agar; TSA: trypticase soya agar; PS:
pseudomonas agar; BA: blood agar.
55
3.3. Histopathology
In 2006 and 2010, outbreaks of gill disease occurred in LT with an average size of 2.5
cm. Even though there was no presence of EPI in these fish, based on the patterns of
necrosis and the lack of other agents associated with mortality in the hatchery history, a
tentative diagnosis of CLO was made. Historically, most of the cases presenting with
visible EPI were reported during winter time; however, clinically affected fish during
2006 were reported in the early spring (March and April). During these two months, gill
samples were collected for histopathological analyses and while characteristic lesions
were present, there was no presence of EPI, nor was there a change in the severity of the
lesions (Figure 3.2).
In 2011, 2012 and 2013 outbreaks of gill disease in BJC occurred in fish with an average
size of 10 cm. The histopathological lesions however were consistent year to year.
Lamellar epithelial hyperplasia, fusion of gill filaments, thrombosis, and epithelial cell
necrosis were observed. In addition the interlamellar spaces were filled with
inflammatory cells and hyperplastic epithelial cells (Figures 3.4, 3.5). Lamellae with a
square profile were also common (Figure 3.6). These lamellae had hypertrophied
epithelium most obvious in their distal half.
Relatively high numbers of EPI were common in gills examined from these years, in
contrast to those from 2006 and 2010. The cells containing bacterial inclusions were
hypertrophied and were most commonly evident in the interlamellar space (Figure 3.6);
however, they were also found at different locations within the gill tissue. Two types of
EPI were observed: slightly basophilic dense homogenous colonies approximately 10 m
56
in size; and slightly granular eosinophilic colonies that were 15 to 20 m in size. Both
types of inclusions were surrounded by halos (Figure 3.7), and when the nucleus was
evident, it was usually pushed toward the periphery of the cell.
EPI were not differentially stained either by PVK or Giemsa techniques; however, the
inclusions stained Gram-negative using the B&H technique (Figure 3.8). In general, EPI
were usually present in affected gills from the initial submission from an outbreak;
however, no inclusions or small numbers of inclusions were observed in gill samples
obtained from subsequent submissions from the same outbreak.
57
A
B
Figure 3.2 Histopathology of lake trout from 2006. A. Fusion of filaments (thick arrows) moderate to severe lamellar epithelial
hyperplasia (4X H&E) B. There is scattered necrosis (thin arrows) but without any visible EPI (40X H&E).
58
Figure 3.3 Histopathology of lake trout gills from 2013, group, A. Areas of gill lamellar fusion (asterisk) (4X H&E); B. Lake trout
gills 2013, group C. Severe interlamellar hyperplasia and fusion. Fusion of whole filaments was also observed occasionally (asterisk)
(4X H&E).
59
*
*
*
Figure 3.4 Histopathology of lake trout gills from 2013, group B. Mucus cell metaplasia (asterisk), scattered single-cell necrosis (thin
arrows), mitotic cells indicating cellular proliferation (thick arrow) and EPI (circle) (40X H&E).
60
Figure 3.5 Histopathology of lake trout gills from 2013, group B (2). Accumulation of eosinophilic granular cells (arrowhead), mucus
cell metaplasia (thin arrow) and necrosis with intraepithelial infiltrates of leukocytes (circle) (40X H&E).
61
Figure 3.6 Histopathology of lake trout gills from 2013 group C. An EPI within the interlamellar epithelium with light basophilic
intracellular inclusion surrounded by a characteristic halo (arrow). Lamellae with a square profile (circle) and necrosis (arrowheads)
can also be observed (60X H&E).
62
Figure 3.7 Histopathology of lake trout gills from 2013, group D. A. Slightly basophilic with densely homogenous content and
periphealized nucleus (thin arrow) ~10 µm EPI (arrow), and necrosis (arrowheads). B. Granular slightly eosinophilic ~20 µm EPI
(arrow) (100X H&E).
63
Figure 3.8 Differential staining of lake trout gills from 2013, group D. EPI were not stained differentially using PVK (A) or Giemsa
(B). However staining on B&H was consistent with Gram negative organisms (C) (60X).
64
Histopathological changes in gills obtained from BJC RT fingerlings during the winter of
2013 (December, January) were semi-quantitatively scored to detect any association with
mortality rates and to detect any variation in the intensity of the lesions during the
progress of this 4-week outbreak (Figures 3.3, 3.4, 3.5, 3.6, 3.7).
Significant differences (p<0.05) were observed for lamellar hyperplasia (LH), necrosis
(N) and the number of EPI among samples obtained at different time points. Conversely,
results for eosinophilic granular cells (EGC) and goblet cells (G) were not significantly
different at any time point (p>0.05) (Table 3.2 and Figure 3.9).
Even though fish obtained in period A did not have any clinical signs there were mild gill
lesions present. However, values for LH in samples of fish obtained in period A were
significantly (p<0.05) smaller than those observed in fish obtained from later time points
(Table 3.2 and Figure 3.9).
Gills from fish obtained before clinical sings were evident (time A) had significantly
(p<0.05) fewer EPI compared to those from fish sampled at times B and D. The numbers
of EPI in fish gills obtained at sampling period C were numerically, but not significantly
(p>0.05), higher than those of fish from time A. Additionally, there were no significant
(p>0.05) differences in the number of EPI found during the last three sampling times.
The severity of branchial necrosis in gills of fish from time A was significantly (p<0.05)
less than those from gills of fish obtained at time C. No other significant (p>0.05)
65
differences were observed among gills of fish examined at times A, B or D. Additionally,
scores for necrosis in gill samples obtained from fish sampled at times when branchial
epithelial necrosis was found (B, C and D) were not significantly (p>0.05) different.
66
Table 3.2 Gill lesion score results from tank R8 sampled between December 3 rd , 2012 and January 18 th 2013.
Date
Lamellar
epithelial
hyperplasia (LH)
Epitheliocystis
inclusions (EPI)
Necrotic cells
(N)
Metaplastic
goblet cells
(GM)
Eosinophilic
granular cells
(EGC)
Mortality on
the sampling
day
December 3
(A)
1.5 ± 0.5c
1.2 ± 0.2b
1.6± 0.3 b
1.7± 0.1
1.5± 0.3
0
January 3
(B)
2.6± 0.3b
1.9 ±0.4a
2.1± 0.5ab
1.7± 0.5
1.9± 0.2
7
January 8
(C)
3.9± 0.2a
1.7± 0.4ab
2.4 ± 0.7a
2.1± 0.8
1.5± 0.4
13
3.5± 0.4a
2.2± 0.4a
1.7± 0.7ab
2.2± 0.8
1.8± 0.4
157
January 18
(D)
Values are means (n=5) ± SE. a, b, c Different superscripts within a column (particular lesion classification) are significantly different
(p<0.05).
67
EPI
Figure 3.9 Gill lesions (mean ± SE) at four different times and mortality of Blue Jay Creek lake trout from tank R8 during 2013. Bars
followed with different superscripts (a, b, c) within dates are significantly different (p<0.05).
68
3.4. PCR and DNA sequencing analysis
3.4.1. Universal bacterial 16S rRNA gene primers
A total of 87 LT were tested from 2010 to 2013 during outbreaks of gill disease (Table
3.3). The PCR protocol UNIRX2 and primers U1 and U1R (Appendix 7.3) were used to
target a ~ 800 base pair sequence of the 16S rRNA gene. All the samples that generated
gel bands were amplified at least twice in separate PCR experiments. A total of 76 bands
were extracted, purified and direct sequenced using the U1 and U1R primers. Two
precautions were taken in order to verify the quality of sequence results. First using the
BLAST- n online software, the forward and reverse sequences were checked individually
in order to verify that they matched the same bacteria. In some cases U1 and U1R
primers yielded different bacterial sequences, this was the case for some 2010 samples
(Appendix 7.4). Second, pairwise alignments between the forward and reverse sequence
was done and checked using DECIPHER online software to detect possible chimera
sequences (http://decipher.cee.wisc.edu/FindChimeras.html). A total of 56 pairwise
consensus between forward and reverse sequences had a 90 % maximum identity (MI) to
a proposed novel organism, CBc (JN968376.1; Toenshoff et al, 2012) that was obtained
from AS with EP in Norway (uploaded in GenBank on April 24, 2012). A multiple
alignment was build using these 56 sequences and those that shared more than 99%
nucleotide identity, were used to obtain a consensus named “BJC-BK”
In the case of bacteria cultured and isolated from agar at either of the two temperatures
(16 ºC or 10 ºC) or in any of the media utilized none of the 16S rDNA sequences
obtained belonged to Burkholderiales (Table 3.1).
69
LCM of samples from 2011, 2012 and 2013 generated 340 bp amplicons that shared 85
%, 86 % and 89 % nucleotide identity with JQ723599.1 (CBc), and 82 %, 82 % and 87 %
with BJK-BK (Table 3.4; Figure 3.11). Samples did not react with Chlamydiales primers.
70
Figure 3.10 Agarose gel (1 %) of 800 bp products generated from lake trout gills using
the PCR protocol UNIRX2 to detect bacterial 16S rDNA . A. 1-5 were from December
3rd (Group A), when there were no clinical signs; 5-10 were from January 3rd (Group B)
at the beginning of the outbreak; 11-20 were taken on January 7th (Group C) when
clinical signs were present and mortality had begun; 21. Positive control, Vagococcus
salmoninarum; 22 and 23. Negative controls, DNA from liver and control template
respectively. B: 1-10 were from January 18th (Group D) when mortality was highest. 11.
Positive control, Vagococcus salmoninarum, 12 and 13. Negative controls, DNA from
liver and control template, respectively.
71
Table 3.3 Number of samples that have 90 % maximum identity with Candidatus
Brachiomona cysticola.
Case number
B126-07
Date
March-April, 2006
Sequence result 90%MI CBc
Not tested (only samples in formalin)
B037-10
February 24, 2010
Negative
B047-10
March 25, 2010
Negative
B028-11
February 11, 2011
Not tested (only samples in formalin)
B034-11
Feb 18, 2011
4out of 6
B042-11
March 3, 2011
8 out of 8
B214-11
December 21, 2011
6 out of 6
B016-12
January, 2012
10 out of 10
B016-12
February, 2012
4 out of 10
B014-13 (A)
December 3, 2012
5 out of 5
B002-13 (B)
January 3, 2013
5 out of 5
B002-13 (C)
January 8, 2013
7 out of 10
B015-13 (D)
January 18, 2013
7 out of 10
72
Table 3.4 Nucleotide similarities between sequences obtained by laser capture microdissection and PCR from DNA directly extracted
from fresh gill tissue of lake trout with epitheliocystis.
Samples
LCM 2011
LCM 2012
LCM 2013
BJK-BK
JQ723599.1 (CBc)
Maximun Identity
2011
100 %
86.4%
85.0 %
82.0 %
85.0 %
JF800717.1 (90.0 %)
100 %
85.0 %
82.0 %
86.0 %
DQ831001.1 (88.0 %)
100 %
87.0 %
89.0 %
EU841510 (90.0 %)
100 %
90.0 %
JQ723599.1 (90.0 %)
100 %
JQ723599.1 (100 %)
2012
2013
BJK-BK
JQ723599.1
(CBc)
73
B
A
Figure 3.11 Tissue sections used for laser capture microdissection. A. EPI before excision (Arrows). B. Spaces are left in excised
areas.
74
3.4.2. 16rRNA Chlamydiales primers
The primer pair 16SIGF and 16SIGR, which generated a ~300bp product, were
problematic. At least two repetitions of each reaction was performed and directly
sequenced. As with 16rDNA primers, the forward and reverse sequences were
individually analysed using the BLAST- n online software, and checked for chimeras
using the DECIPHER software. Most of the sequences obtained using the 16SIGF and
16SIGR primers yielded different Chlamydiales, or in some cases only 16SIGR was able
to generate a quality sequence (Table 3.5 and Figure 3.12). In addition, samples that
yielded a ~300 bp Chlamydiales product were also run with the 806R/16SIGF and
16SB1/16SIGF primer pairs in order to obtain ~800 bp and ~1500 bp products,
respectively. Neither of those primer pairs were able to amplify any product (Appendix
7.2).
A new set of primers, FOR2 and REV2, that amplified a ~270 bp of Chlamydiales
product were therefore tested (Ossewaarde and Meijer, 1999). With this set of primers,
strong bands that could easily be sequenced were obtained, producing much more
consistent sequencing results (Table 3.5; Appendix 7.3). LT samples analyzed with the
protocol CLORX4TD using primers FOR2/REV2 (Tables 3.5; Appendix 7.3) were
amplified at least twice in independent PCR experiments. A total of 34 out of 84 samples
between years 2010 and 2013 produced sufficiently strong bands to allowed purification
and direct sequencing. As in previous PCR reactions, the forward and reverse sequences
where individually checked using BLAST-n and DECIPHER online software to detect
chimeras. Pairwise alignments of 34 individual sequences of ~270 bp were generated
75
from the forward and reverse primer. The resulting pairwise alignments were analyzed by
BLAST-n. and a total of 7 sequences were matches for Candidatus Piscichlamydia sp. E4
(JX470313.1) with a MI of ~94 %. A multiple alignment was made out of these 7
sequences and the consensus obtained was named “CHLA-BJC2”. There were matches
for at least other 9 different Chlamydiales sequences in the Genbank nucleotide database
however, significantly less often than Candidatus Piscichlamydia salmonis E4 (Table
3.5).
76
Table 3.5 Sequences (270 bp) identified in lake trout during epitheliocystis outbreaks using 16S Chlamydiales
primers.
Positives/
total of
samples
Nucleotide
data base
number
Maximun
identity
2/8
JQ480300.1
87%
Chlamydiales in African
catfish (Clarias
gariepinus)
1/8
FJ160740.1
85 %
1/8
GU968541.1
1/8
1/8
1/8
Year of
sampling
Organism
Primer pairs
Notes
2010
Everett et al.,
1999
Sequenced named
CHLA-BJC1
eye infection in sheep
2010
Everett et al.,
1999
88 %
Chlamydia from sea bird
feces
2010
Everett et al.,
1999
Only 16SIGR gave a
product
NR_074972.1
82 %
Parachlamydia
acanthamoebae
2010
Everett et al.,
1999
Only 16SIGR gave a
product
JN616113.1
86 %
Lava cave bacterial
diversity
2010
Everett et al.,
1999
Only 16SIGR gave a
product
NR_074972.1
88 %
Parachlamydia
acanthamoebae
2011
Ossewarde and
Meijer, 1999
77
Table 3.5 (Continued from page 75) Sequences (270 bp) identified in lake trout during epitheliocystis outbreaks
using 16S Chlamydiales primers.
Positives/
total of
samples
2/8
1/8
1/8
1/8
1/8
6/12
1/12
Nucleotide
data base
number
Maximun
identity
FM176910
90 %
JX470313.1
FJ817587.1
GU918153.1
HQ702416.1
JX470313.1
HQ702416.1
Organism
Year of
sampling
Primer pairs
Uncultured neochlamydia
2011
Ossewarde and
Meijer, 1999
93 %
Candidatus
Piscichlamydia sp. E4
2011
Ossewarde and
Meijer, 1999
87 %
Uncultured Chlamydiae
bacterium clone UMAC4
2012
Ossewarde and
Meijer, 1999
89 %
Uncultured bacterium
2012
Ossewarde and
Meijer, 1999
93 %
Uncultured
Parachlamydiaceae
bacterium clone WT41
2012
Ossewarde and
Meijer, 1999
94 %
Candidatus
Piscichlamydia sp. E4
2013
Ossewarde and
Meijer, 1999
100 %
Uncultured
Parachlamydiaceae
bacterium clone WT41
2013
Ossewarde and
Meijer, 1999
78
Notes
Sequenced named
CHLA-BJC2
Sequenced named
CHLA-BJC2
Figure 3.12 Agarose gel (2 %) of 270 bp products generated from lake trout gills using
the PCR protocol CLORX4T. A. 1-5. December 3rd (Group A), when there were no
clinical signs; 5-10. January 3rd (Group B) at the beginning of the outbreak; 11-20.
January 8th (Group C) when clinical signs were present and mortality had begun; 21.
Positive control Candidatus Piscichlamydia salmonis; 22 and 23. Negative controls,
DNA from liver and control template respectively. B: Samples from 1 to 10 were taken
during peak mortality, on January 18th (Group D); 11. Positive control Candidatus
Piscichlamydia salmonis. 12 and 13. Negative controls, DNA from liver and control
template, respectively.
79
3.4.3. EEDV primers
A total of 84 LT gills collected from 2010 to 2013 were examined using the 339F/340R
primer pair, and the PCR protocol EEDVRX1 to screen for the presence of EEDV. No
PCR product was generated using these primers (data not shown).
3.5. Molecular and phylogenetic analyses
Samples on which 16S bacterial universal PCR primers were previously used to amplify
a 800 bp product were analysed again using primers that generated a full-length (1500
bp) product to provide more sequence data for molecular phylogenetic analysis.
Molecular phylogenetic analysis based on the Jukes Cantor model revealed that the
bacterium BJC-BK was genetically related to members of the order Burkholderiales BJCBK shares at least 90 % nucleotide identity with two other organisms that have been
associated with PGI and EPI in AS from Europe, JN807444 and CBc. These two agents
and BJC-BK are proposed to be in the same family (Figure 3.13).
80
BJC-BK
Figure 3.13 Order Burkholderiales phylogenetic tree inferred by using the neighbour
joining method, Jukes Cantor model of the full-length16SrRNA sequences of known
Burkholderiales members and BJC-BK. The outlier-group is Piscirickettsia salmonis
81
A consensus sequence of 277 bp was obtained after using the Chlamydiales primers
FOR2 and REV2. Reliable phylogenetic analyses are difficult to perform in sequences
less than ~300 nucleotides in length (Corsaro et al., 2003). However in the case of
Chlamydiales, it has been proposed that the first 300 bp of the 5’ to 3’ 16S segment has
enough variability between species to allow taxonomical classification (Everett et al.,
1999; Meijer and Ossewarde 2002). Molecular phylogenetic analysis based on the Jukes
Cantor model revealed that the bacterium CHLA-BJC2 was genetically related to
members of the order Chlamydiales (Figure 3.14). CHLA-BJC shares at least 93 %
nucleotide identity with Candidatus Piscichlamydia sp. E4 (JX470313.1).
82
Figure 3.14 Order Chlamydiales phylogenetic tree inferred by using the neighbor joining
method, Jukes Cantor model of the partial-length (~300 bp) of 16SrRNA sequences of
known Chlamydiales members and CHLA-BJC2. The outlier-group is Piscirickettsia
salmonis.
83
3.6. Immunohistochemistry
None of the EPI from LT were positive with IHC using the monoclonal antibodies raised
against Chlamydia trachomatis LPS (Figure 3.15).
84
Figure 3.15 Immunohistochemistry of lake trout gills from 2013, group D: A. EPI
(arrowhead) (40X H&E): B. Same microcolony as in A; no immunoreactivity using the
Chlamydia trachomatis MoAb 163 (arrowhead) (40X); C. sheep placenta infected with
Chlamydia abortus as a positive control (20X).
85
3.1. Transmission electron microscopy
Transmission electron microscopy was performed on formalin fixed paraffin embedded
gill tissues from clinically affected fish from which EPI had been detected using light
microscopy. EPI in glutaraldehyde fixed tissue were not found. No differential staining of
the inclusion was found in semi-thin sections (Figure 3.16 A and B). The organisms were
pleomorphic and varied from coccoid structures (~ 0.5 to 1.0 m) to long rods (2.5 m)
and apparently divide by budding (Figure 3.17. 3.18, 3.19). In some cases contained
central electron-dense material (nucleoids) and electroluscent vesicles (3.20). A
comparison between the BJC LT EP agent and descriptions from another study is
summarized in Table 3.6.
86
Table 3.6 Comparison between organisms identified by electron microscopy in previous reports and the EPI
organism described in the present study.
Candidatus
Clavochlamydia
salmonicola
Candidatus
Piscichlamydia
salmonis
Epitheliocystis
agent in
Artic Charr
Candidatus
Brachiomona
cysticola
BJC-BK
epitheliocystis
agent
Inclusion size
15 m
10 m
10 m
10 m
10 m
Membrane bound
Yes
Yes
Yes
Yes
Yes
Organism shape
EB- IB- RB-HT
(0.5 m)
EB- IB- RB
(0.3-0.7 m)
RB
(0.5-2.5)
Rounded to elongated
forms, approximately
0.2–0.4 (diameter)x<2
m (length)
Rounded to
elongated forms,
~0.2–0.4
(diameter)x<2 m
(length)
Matrix
Not osmiophilic, IB
and RB distributed
perifelically within
the inclusion
Moderately
osmiophilic,
fibrillar.
Moderately
osmiophilic,
fibrillar
Osmiophilic
Osmiophilic
Taxonomic
classification
Chlamydiales, family
Chlamydiaceae
Chlamydiales
Chlamydiales
Beta-proteobacteria,
Burkholderiales
Probably a Bproteobacteria
Burkholderiales
Criteria
EB: elementary bodies; IB: intermediary bodies; Reticular bodies; HT: Head tail bodies
87
Figure 3.16 Transmission electron microscopy of a lake trout gill from a fish affected by epitheliocystis. A. Semi-thin section with
one EPI at the base of a lamella (dotted circle); B. EPI (arrow) from the semi-thin section identified using transmission electron
microscopy.
88
Figure 3.17 Transmission electron microscopy of lake trout gills affected by epitheliocystis. An EPI (arrow) is localized in the
epithelium between two lamella with pillar cell channels (arrowhead). The thin arrow indicates the base of the interlamellar epithelial
zone.
89
Figure 3.18 Transmission electron microscopy of lake trout gill affected by epitheliocystis (2). EPI with polymorphic bacteria present
in the center of the cell surrounded by a “halo”
90
Figure 3.19 Transmission electron microscopy of lake trout gill affected by epitheliocystis (3). Same EPI from figure 3.17 with
polymorphic bacteria dividing by budding (arrowheads).
91
*
*
Figure 3.20 Transmission electron microscopy of lake trout gill affected by epitheliocystis (4). EPI bacteria with double membranes
(arrows), nucleoids (arrowheads) and electroluscent vesicles (asterisk).
92
4. DISCUSSION
Lake trout are raised for release by the OMNR at BJC Fish Culture Station, Manitoulin
Island, ON. This facility has reported yearly epizootics of EP during the winter associated
with mortalities. Based on previous studies regarding EP in different fish species and the
characteristic lesions observed during histopathologic examination, it was thought that
the causative agent of EP in LT at BJC might be a bacterium that belongs to the order
Chlamydiales. However, results based on PCR, EM and LCM indicate that EP in LT at
the BJC Fish Culture Station is associated with a novel ß-proteabacteria that belongs to
the order Burkholderiales. On the other hand, Chlamydiales were detected using PCR in
both clinically healthy and EP-affected fish, but not in EPI retrieved by LCM. Therefore,
the first hypothesis proposed in this thesis should be rejected. This is the first study to
associate a member of the order Burkholderiales with cases of EP in any fresh water fish
and the second in any fish species.
Cases of EP in LT in the BJC culture station generally occur in the winter (DecemberFebruary). These outbreaks have been temporally related to heavy rain events causing
melting of snow and floods. The water temperature in the holdings facilities is ~ 8ºC
during that time of the year. Underground water becomes turbid due to the effect of
runoffs, resulting in an increase in suspended solids (SS) in the water, and soil
accumulates at the bottom of the tanks. Studies in Norway have shown that EP caused by
93
CBc or/and CPs in AS, generally occurs during the fall following spring seawater transfer
(Steinum et al., 2009), while EP in BT caused by CPs and CCc in Switzerland, occurs
during peak water temperatures in the summer (Schmidt-Posthaus et al., 2011). Wild BT
did not show any clinical signs associated with the presence of EP agents during the
summer time, which differed from Norwegian AS. It is evident that EP agents can be
present at any time during the year; however, mortality seems to be related with EP in
colder water temperatures which is the case of Norwegian AS and BJC LT.
Exposure to increased levels of SS (suspended solids) during floods or heavy rains at BJC
may play a role in LT EP and associated mortality events. It has been demonstrated that
exposure to prevailing levels of inert kaolin particles has little direct effect on gill health
(Goldes et al., 1988). However, SS produce alterations in water quality reducing
dissolved oxygen and increasing temperature (Oschwald, 1972; Appleby and Scarratt,
1989; Marcus et al., 1990). Additionally, fish are able to tolerate short episodes of
extremely high levels of suspended sediment by exuding protective mucus on the skin
and gills. This mucus traps and continually removes trapped particles but comes at a
metabolic cost, which may place the fish under stress (Persaud and Jaagumagi 1995). It
has been demonstrated in coho salmon and rainbow trout that even though they survive
exposure to high concentrations of SS, they may undergo sub-lethal physiological stress
that reduces their physiological performance capacity (Redding et al., 1987).
A decrease in water temperature during the fall and winter season, and increased
exposure to SS, are conditions which are favourable for an increase in the concentration
94
of potentially pathogenic organisms such as /bacteria, protozoans, and even amoeba.
These conditions may allow greater contact of these agents with the gills of
physiologically stressed, potentially immunosuppressed, fish. Once EP agents have
invaded, environmental or immunological impairment may allow for bacterial
proliferation. Once EP agents have colonized the tissue, virulence determinants, such as
those that allow proliferation and persistence, may be expressed. Further studies of the
pathogenenesis are required before definitive conclusions can be drawn, and
experimental transmission would facilitate this effort. The exact cause of the increased
mortalities that occur during wintertime in BJC LT is unclear. It is possible that EP might
be part of a multifactorial disease complex similar to PGI in AS, and there may be more
than one agent that contributes to the clinical presentation
(Nylund et al., 1998;
Kvellestad et al., 2005; Steinum et al., 2009). EPI were the most characteristic
histopathological lesion observed during outbreaks; however, there is insufficient
evidence to conclude that this agent was primarily responsible for the mortality in the
BJC LT population.
PCR using primers designed to bacterial 16S rRNA gene sequences demonstrated that a
ß-proteobacterium from the Order Burkholderiales (BJC-BK) is consistently present in all
the outbreaks studied from 2011 to 2013 in BJC LT. This is supported by LCM, which
demonstrated that a ß-proteobacterium belonging to the order Burkholderiales was
identified in intracellular microcolonies in BJC LT cases from 2011, 2012 and 2013.
However, PCR sequences of DNA extracted from fresh gill tissue and LCM derived
sequences of DNA extracted from FFPE tissue, only shared ~82 % nucleotide identity
95
with BJC-BK in year 2011 and 2012 and 87 % in year 2013; therefore, this is not
conclusive. One explanation for the relatively low nucleotide identity may be that
sequences resulting from LCM using FFPE tissue, resulted in low-complexity sequences
(Wootton and Federhen, 1996), leading to mismatches during alignment with the
sequences obtained from fresh gill tissue.
Bacteria from the order Burkholderiales are associated with respiratory disease in birds,
such as bordetellosis in wild and domestic birds, and Burkholderia cepacia in humans
which is associated with pneumonia in immunocompromised individuals with lung
disease (Mahenthiralingam et al., 2007; Harrintong et al., 2009). In the case of fish, a
novel bacterium, CBc, from the order Burkholderiales, has been identified as the
causative agent of EP associated with PGI in Norway and Ireland (Toenshoff et al., 2012;
Mitchell et al., 2013). The BJC-BK, bacteria associated with EP in BJC LT is a yet
unidentifed agent that shares 90 % nucleotide identity with CBc.
Using Chlamydiales 16S rRNA gene primers, several chlamydial sequences were
obtained from fresh gill tissue during different years (Table 2.1). There is evidence that
CPs is certainly associated with the examined gill tissue, although it appears not likely to
be the agent of EPI in these cases. This conclusion is also supported by LM results; the
EPI were not stained by PVK and Giemsa, i.e. stains that are usually used to identify
Chlamydiales in intracellular inclusions of birds, humans and fish (Elder and Brwon,
1999; Draghi et al., 2007). In addition, monoclonal antibodies raised against Chlamydia
trachomatis LPS did not react with EPI (Figure 3.15). These results also support the
96
findings of Toenshoff et al. (2012) who examined gills of AS with EP and demonstrated
that both CPs and CBc are present using FISH; however, only CBc were within
intracellular colonies, while CPs appeared to be diffusely widespread throughout the gill
tissue.
The predominant histopathological pattern in LT gills affected by EP is proliferative,
similar to that of PGI described in farmed AS, where lamellar hyperplasia and fusion are
most prominent but inflammation and necrosis are also notable (Kvellestad et al., 2005;
Steinum et al., 2009; 2010). Circulatory disturbances such as thrombosis or death of pillar
cells, which are part of the pathological changes associated with PGI, were also observed
in BJC LT, but they were not consistently present in BJC EP outbreaks.
Morphological similarities between cases of EP caused by CPs and CBc, include the
dense homogeneous basophilic content and the size (10 µm) of the EPI;these
characteristics and the host reaction associated with these two agents are very similar to
BJC EPI (Draghi et al., 2004; 2010; Schmidt-Posthaus et al., 2011; Toenshoff, et al
2012). On the other hand there are some differences that are observed in LM, for example
BJC EPI are Gimenez negative, while CPs stain positive with this stain (Draghi et al.,
2010). Further, while CBc and BJC EPI are both Gram negative, a slightly difference in
the shape of the colonies can be observed; CBc have rounded, well-defined edges, while
BJC EPI are more irregular. Still, EPI size, shape, staining and tissue host reaction seem
to be quite similar between CBc and the BJC EPI agent. The two different types of BJC
97
EPI (~10µ and ~20µ) may indicate different stages of inclusion development, but the
~20µ EPI were not found using TEM and so the inclusions could not be compared.
EP cases at BJC also have histological similarities with EP caused by Chlamydiales
infection; however, the ultrastructural cell morphologies observed within the intracellular
inclusions were not consistent with a chlamydial life cycle. The bacteria in BJC LT EPI
are pleomorphic, from coccoid to rod-shaped, and appear to be closer in shape to the ßproteobacteria described by Toenshoff et al. (2012) than to Chlamydiales EB or RB, but
taxonomic conclusions cannot be made based on ultrastructure alone.
From previous subjective evaluation of BJC cases, it was proposed that the numbers of
EPI appeared to be higher at the beginning of the outbreak but were decreasing when the
mortalities peaked. Therefore, by the time that most samples are sent for laboratory
examination, necrosis and lamellar hyperplasia, but not EPI, were the most outstanding
lesions.
To investigate this, an objective evaluation of histological lesions in LT from BJC was
performed. The scoring system was designed to include the most common pathological
lesions observed during the disease outbreaks over the past years and was applied to the
2013 gill samples. Samples were taken at four different time periods and were evaluated
using a semi-quantitative scoring system. All of the characteristic lesions observed in LT
during the EP outbreak were present to a mild degree in fish with no clinical signs at the
first sample time. A survey conducted on 48 species of ocean-caught fishes from British
Columbia, Canada indicated that 25 species had EPI even though the fish had no clinical
98
signs (Kent et al., 1998). It is therefore clear, therefore, that a subgroup of a population
can be infected with an EP agent without showing clinical signs. In the present study,
gills in period A had EPI present, indicating either an early winter outbreak, or as in the
case of wild fish in British Columbia, EP agents may be present in a population during
the entire year and are not always associated with mortality events. This idea is supported
by a recent study using qPCR performed in Norwegian and Irish AS (Mitchell et al.,
2013), where the EPI agent CBc was present in clinically healthy fish, but also increased
in fish during an outbreak.
The number of EPI in fish with clinical signs of infection (periods B, C and D) were
significantly higher than those observed in fish sampled in period A (no clinical signs);
consequently, the second hypothesis proposed in this thesis might be true. However, there
was not a significant difference in EPI number from period B to D, when mortality
reached the peak on period D. Further research in LT, using a qPCR would help better
answer this question.
There was a significant difference in necrosis between the fish with no clinical signs in
period A and the fish sampled in period C, when mortalities were just starting to rise.
Contradictorily, there was no significant difference in necrosis between periods C and D
where the mortalities are reaching the peak and clinically healthy fish in period A. The
sample size examined at each time may have been too small (five fish) to detect any
differences, additionally samples from the population after mortality has ceased were not
available at the time of analyses.
99
The lesion that was most closely related to the severity of clinical signs and mortality was
LH. A proliferative cell response that resulted in the gradual destruction of the functional
integrity of the gills has been suggested as the cause of massive mortalities in cultured
Yellowtail (Seriola mazatlana) with EP from Ecuador (Venizelos and Benetti, 1996).
However, there are also cases where there is no proliferative host response associated
with EPI, for example in Australian bass (Macquaria novemaculeata) and bartail flathead
(Platycephalus indicus), where mortalities reached up to 100 % (Nowak and LaPatra,
2006).
For this longitudinal study, samples of gill tissue for PCR analysis were collected. BJCBK was present during all four sampling periods and this agrees with the histological
presence of EPI during all four periods. During period C the bands produced by 16S PCR
are more prominent, which provides a tantalizing possibility of higher bacterial
concentrations, since loading of gels was standardised. However a specific and
quantitative PCR (qPCR) should be developed to evaluate bacterial concentration and
prevalence associated with periods of health and episodes of gill disease (Fredricks and
Relman, 1996).
Antimicrobials used to date to treat cases of EP in LT (e.g. tribrissen and oxytetracycline)
have not proven effective in limiting the impact of outbreaks (not shown). Since the
pathogenesis of the disease is not conclusively understood at present, dealing with the
environmental stress factors such us suspended solids would be perhaps a potential
100
solution in this particular case. Setting of a silt fence, sediment traps and a membrane
filtration system along with UV treatment of the water should be considered, although the
expenses required for these would be considerable.
Currently, there is enough evidence demonstrating that EP can be caused by bacteria
other than Chlamydiales (Toenshoff et al., 2012; Mitchell et al., 2013; Mendoza et al.,
2013). Consequently, the term CLO when referring to histology lesions should be used
carefully to avoid misinterpretations about the true etiology of the “inclusion” and of the
disease. Additionally, even though EPI have been associated with mortality events in
several cases (Nowak and LaPatra, 2006), there is insufficient evidence to correlate
mortalities with a Chlamydiales etiology. However, definitive correlation has been shown
between mortalities and EP caused by Burkholderiales organisms (Toenshoff et al., 2012;
Mitchell et al., 2013).
The study of the outbreaks of EP at BJC in 2011, 2012 and 2013 all display three
common features: the presence of BJC-BK, EPI and a mortality rate of ~40% each year.
The association between the presence of EPI and mortality events in BJC is not clear.
LCM has shown that Burkholderiales and not Chlmaydiales are more likely the cause of
EP in BJC; however, LCM sequences share only 86% nucleotide identity with BJC-BK.
The direct association between BJC-BK bacteria and EPI will be tested using in situ
hybridization in the near future. Additionally, the concentration of BJC-BK bacteria
associated with yearly disease outbreaks will be checked using a qPCR, based on the
work described here for this new bacterium. Clinically healthy fish and water will also be
101
tested using the qPCR in order to determine bacterial concentrations associated with the
fish and environment (Fredricks and Relman, 1996).
Further studies of the pathogenesis and epidemiology of EP are needed. While EP agents
have not yet been cultured, molecular methods such as PCR and in situ hybridization
should be used to probe the relationship between EPI and an aetiological agent. Culture
media conditions might be hard to replicate due the intracellular nature of EP agents in
general. It has been demonstrated that free-living Acanthamoeba spp are hosts for
numerous pathogenic bacteria such as Burkholderia cepacia, Legionella pneumophila,
Mycobacterium avium and several Chlamydiales strains (Landers et al., 2000; Kilvington
and Price, 1990; Steinert et al., 1998; Corsaro et al., 2009). Consequently amoeba coculture techniques should be considered as potential methods for isolation of EP agents
beside the other in vitro culture techniques (cell and media culture).
102
5. CONCLUSIONS
1. Based on electron microscopy, cell morphologies observed within the cytoplasmic
inclusions in the present study do not appear to be consistent with a chlamydial life cycle.
2. Based on polymerase chain reaction results, Chlamydia-like organisms are associated
with the tissues of infected fish. Their presence as a normal microbiota or part of the
disease complex needs to be resolved.
3. The agent most consistently associated with epitheliocystis in Blue Jay Creek lake
trout is a novel bacterium that would belong to the same family as Candidatus
Brachiomona cysticolas. The clone BJC-BK has been uploaded to NCBI and a name for
this new organism will be proposed.
4. Epitheliocystis inclusions and lamella hyperplasia were positively associated with
increased mortality in Blue Jay Creek lake trout.
103
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7. APPENDICES
7.1 EP-PM Protocol 1
Important: Prepare the materials at least a couple of hours before sampling to decrease
the amount of gill decomposition during processing. Please see where to get things and
where to store samples in the last section.
1.
Materials
a)
Histology cassettes
b)
Tubes 1.5 microcentrifuge tubes with RNA later
c)
Empty 1.5 microcentrifuge tubes
d)
100 ml yellow cap formalin plastic container
e)
1 300 ml formalin plastic container
f)
Plastic bag for freeze tissue (whirlpool)
g)
Carcass disposal bag (Biohazard)
h)
Gloves and lab coat
i)
Dissection tools (scissor, blade and forceps)
j)
Flame
k)
Box for 1.5 ml microcentrifuge tube
*Before: start label all the materials with the following information (bags, box and tubes)
1) B# (the following number in the lab case book). Please give a new B# every time that
fish are sampled (less confusing for histology room people)
119
2) Place of sampling (BJC)
3) Specie (LT)
4) Date (tubes do not need the date)
5) Name of the researcher
Shorter form of common names for labelling sampling material
Species:
RT: rainbow trout
AC: Arctic charr
LT: lake trout
BT: brown trout
KO: koi
Places:
Alma
BJC (Blue Jay Creek)
CAF (Central Animal Facility)
2. Methods
Gills and Skin
1)
Collect the number of fish needed per tank (recently euthanized) and place them
on ice immediately
2)
First arch: place half in microcentrifuge tube and half in histo cassettes.
3)
Second arch: place half in plastic bag (to be frozen) and half in formalin (100 ml)
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4)
Third arch will be placed in RNA later (cut it in three pieces). Do not overload the
Tube; the cap should be able to close well
5)
Fourth arch: save entirely in the plastic bag (to be frozen, same bag)
6)
Proceed the same way with the gills in the other side of the fish
7)
Before opening the carcass, cut three pieces of dorsal skin and place one in 100ml
formalin, one in plastic bag and the last one in the cassette
8)
Place the cassette in the 500 ml formalin flask
9)
Close microcentrifuge tubes and place them on ice
Internal organs
1)
Important organs to collect: heart, liver, spleen, head and tail kidney
2)
Each of these organs will be placed in plastic bag and 100ml formalin flask
3)
For the 100 ml formalin flask just 2 cm squares of tissues are needed
4)
The rest of the organ that are not placed in formalin should be
saved in plastic bags
5)
Put fish carcass in a biohazard bag for posterior disposal in PM room 1st
Floor of Pathobiology building
3. Disposals and disinfection
1) Between each fish the dissections tools should be placed in 70 % ethanol and
flamed
2) The fish left over are placed in a plastic bag and taken down to the PM
mammalian rooms for destruction.
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3) Clean and disinfect tools with detergent and brush the PM table surface with the
same detergent. Rinse with water
4) Gloves should be disposed in biohazard bags
4. Where to find material to prepare sampling and some recommendations
1)
1.5 microcentrifuge tubes : PCR room. Take a flask containin autoclave tubes and
label it with your name. Keep it on your bench until finish it. Please remember to
not bring it back to the PCR room
2)
Boxes for 1.5 microcentrifuge tube storage are at Elena’s bench under knee
3)
Biohazard and whirlpool (for freezing tissue) bags PM room white boxes under
scope table
4)
Histology cassettes: Trimming room first floor
5)
Formalin flasks: Trimming room first floor
6)
RNA later: PM room cabinet (Usually each of us prepare our own, you could find
how to prepare it in the lab protocols files in droopbox)
7)
Register where your tissues samples were stored
a)
Box with tissues in tubes: FPL -80 ºC 4th floor freezer room
b)
Whirl pack bags: -20 ºC Basement (chest freezer). Recommendation: get a small
cardboard box and keep all your tissues in bags there. It will be easy to find them
in the future
c)
Label everything, (things get lost easily in the freezers)
8)
Finally the best thing to do is prepare a plastic container with all the necessary
things for your future samplings (including markers and pencils)
122
7.2 Primers trouble shooting
Most of the EP studies in fish had used the 16S signature sequence primers set from
Everett et al.(1999). However, in this study, these primers amplify Oncorhynchus mykiss
immunoglobulin heavy chain constant region (ighd) partial gene (Figure below), even
though there is more than 10 miss matches between the primer and the amplified
sequence. This is the first time that molecular methods have been used to identify the EP
agent in LT; this may be the reason why this problem has not been reported in fish
before.
123
7.3 PCR protocols
PCR protocol CLORX1
Step
Initial denaturation
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Final extension
Primer: 16SIGF + 16SIGR
Temperature
94◦C
94◦C
66◦C
72◦C
94◦C
65◦C
72◦C
94◦C
64◦C
72◦C
94◦C
63◦C
72◦C
94◦C
62◦C
72◦C
94◦C
61◦C
72◦C
72◦C
Time
15’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
45’’
7’
PCR protocol CLORX2
Step
Initial denaturation
Denaturation
annealing
Extension
Final extension
Primers: 16SIGF+806R
Temperature
94◦C
94◦C
55◦C
72◦C
72◦C
Time
15’
30’’
45’
45’
7
124
X3
X3
X3
X3
X3
X 25 cycles
X 40 cycles
PCR protocol CLORX3
Step
Initial denaturation
Denaturation
Annealing
Extension
Final extension
Primers: 16SIGF+ 16SB1
Temperature
94◦C
94◦C
58◦C
72◦C
72◦C
Time
15’
40’’
40”
45”
7’
PCR protocol CLORX4
Step
Initial denaturation
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Denaturation
Annealing
Extension
Final extension
Primers: FOR2 + REV2
Temperature
95◦C
95◦C
62◦C
72◦C
95◦C
61◦C
72◦C
95◦C
60◦C
72◦C
95◦C
59◦C
72◦C
95◦C
58◦C
72◦C
95◦C
57◦C
72◦C
72◦C
Time
10’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60’’
60”
60’’
10’
125
X 40 cycles
X2
X2
X2
X2
X2
X 40 cycles
Protocol UNIRX1
Step
Initial Denaturation
Denaturation
Annealing
Extension
Final extension
Primers: RW1+DG74
Temperature
95◦C
96◦C
56◦C
72◦C
72◦C
Time
5’
30’’
60’’
60’’
10’
Protocol UNIRX2
Step
Initial denaturation
Denaturation
Annealing
Extension
Final extension
Primers: U1+U1R
Temperature
94◦C
96◦C
60◦C
72◦C
72◦C
Time
3’
15”
1.5’
2’
5’
Protocol UNIRX3
Step
Initial denaturation
Denaturation
Annealing
Extension
Final extension
Primers: 27f+U1492R
Temperature
95◦C
96◦C
56◦C
72◦C
72◦C
Time
5’
30”
60”
60”
10”
126
X 30 cycles
X 30 cycles
X 30 cycles
7.4 Sequencing results from samples of 2010 using 16S universal
bacteria primers.
Sample
Nucleotide data base
number
Organism
Maxim
identity
1
1UR: FN421535.1
Uncultured bacterium partial 16S
rRNAgene, clone 14
81 %
2
1UR: KC246342.1
Uncultured beta proteobacterium
87 %
3
1UR: AJ295499.1
Uncultured rape rhizosphere bacterium
83 %
4
1UR: AB698045.1
Uncultured bacterium gene for 16S
rRNA, partial sequence, clone: CN42
98 %
1UR: AJ295499.1
Uncultured rape rhizosphere bacterium
wr0041 partial 16S rRNA gene
80 %
1U: JF697452.1
Uncultured bacterium clone reservoir71 16S ribosomal RNA gene, partial
sequence
5
83 %
1UR: KC246148.1
Uncultured beta proteobacterium clone
XSLA094 16S ribosomal RNA gene,
partial sequence
94 %
U1: GU776500.1
Uncultured bacterium clone
F776O8Q01BHO06 16S ribosomal
RNA gene, partial sequence
73 %
1UR: HM557268.1
Uncultured bacterium clone BICP1313
16S ribosomal RNA gene, partial
sequence
91 %
1U: AB717035.1
Uncultured Pelomonas sp. gene for
16S ribosomal RNA, partial sequence,
clone
88 %
6
7
127
(continued from page 120). Sequencing results from samples of 2010 using 16S universal
bacteria primers.
Sample
Nucleotide data base
number
Organism
Maxim
identity
U1R: AB630695.1
Uncultured bacterium gene for 16S
ribosomal RNA, partial sequence,
clone
86 %
U1: CU920026.1
Uncultured Betaproteobacteria
bacterium 16S rRNA gene from clone
QEDT3BG07
82 %
U1R: AY649345.1
Uncultured bacterium DGGE band m
16S ribosomal RNA gene, partial
sequence
87 %
U1: JX020775.1
Uncultured bacterium clone
L1_003a_A03 16S ribosomal RNA
gene, partial sequence
94 %
U1R: JF809158.1
Uncultured bacterium clone CPf1-G6
16S ribosomal RNA gene, partial
sequence
87 %
U1: KC299368.1
Uncultured bacterium clone 1602 16S
ribosomal RNA gene, partial
sequence.
76 %
Uncultured Brevundimonas sp. clone
LPB-15 16S ribosomal RNA gene,
partial sequence
97 %
8
9
10
U1R: JX855294.1
11
128
(continued from page 121). Sequencing results from samples of 2010 using 16S
universal bacteria primers.
Sample
Nucleotide data base
number
Organism
Maxim
identity
U1R: JF681707.1
Uncultured bacterium clone RMWS15
16S ribosomal RNA gene, partial
sequence
87 %
U1: JX948664.1
Uncultured bacterium clone 4II.46 16S
ribosomal RNA gene, partial sequence
80 %
U1R : JX020775.1
Uncultured bacterium isolate DGGE
gel band 2a 16S ribosomal RNA gene,
partial sequence
95 %
U1: HM599591.1
Uncultured bacterium clone 66111b3
16S ribosomal RNA gene, partial
sequence
84 %
U1: JF412882.1
Uncultured Phenylobacterium sp.
82 %
U1R: HQ910791.1
Uncultured bacterium clone EW_53
16S ribosomal RNA gene, partial
sequence
91 %
12
13
14
129